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UNEXPECTED WAYS YOU ARE LOSING MONEY AT YOUR LAB

Are you a laboratory owner? So what is the core benefit you want when you are running a lab business? Isn’t the answer ‘money’? Revenue and Profits are the core reason why any business is run.

With this post we’ll share unexpected ways in which you are losing money at your lab and things you can do to plug these negative forces.

Website

Well, to be frank a website today has become a basic necessity. It has become essential to have an online presence in the times when almost everyone in the city owns a smartphone. More and more clients are running through online directories, reviews, ratings, answers to shortlist the perfect laboratory. So why should you stay behind? Be present where the clients are present.

A very basic website should work just fine if it can provide the correct information about you and satisfy the client. SEO’s and PPC’s are an add-on advantage & the right keywords can take you places.

Referral Tie Ups

Today more than ever labs need to have referral tie ups with chemist, lab technicians, analyst, doctors, practitioners, health service providers, et al. Overall service is the key. Take for instance, a patient walks in to a doctor’s clinic and the doctor writes some tests, the doctor is now is in a position of influence and can refer the patient to your lab to get the tests done. What does it mean for you? Consistent revenues!

Provide all Services under one roof

Yes, being a specialist in a particular field is cool. But would it be too bad if you become a one stop shop for all laboratory needs? Definitely NOT. Expand your services and provide all solutions to your clients under one roof. The obvious answer here is becoming a partner of choice for your clients A place where the client knows he’ll find all solutions.

Take Control

Yes, you cannot be in two places at one! But being the owner of the laboratory you need to be in control. You need to know what is happening in Branch A or B or rather at Counter No. 3 of Branch C of your labs. Operations form the backbone of any laboratory and those should always be at your fingertips when required. A good way to do that is automation. Automation helps you take control of things and know the processes which are happening in the background.

Inventory Mismanagement

There is nothing worse than losing a customer or a client because some inventory was out of stock. Train your employees in such a way that they keep logs of the consumables and keep note when things go out of stock. A great option in such cases is have an inventory management software. Not only does it reduce reliance on manual counting and log books, it also automates inventory tracking and reordering.

Reference

https://caredatainfo.com/blog/unexpected-ways-losing-money-lab

 

Posted by Akinbuli Opeyemi,

www.aasnig.com, opeyemi@aasnig.com

07084594004, 08068129603

How to make your test sieve last longer?

Test sieves will perform accurately for many years with proper care and maintenance. To make your test sieves last longer, you need to carry out cleaning carefully and use the following tips provided below.

Cleaning Tips:

  • When the particles are lodged in the woven wire mesh, they should be cleaned out as soon as possible. They can enlarge the mesh holes, in turn, affecting the accuracy of the test results.

  • It is recommended to use a special sieve brush to clean the mesh using a gentle circular motion from the underside. Be aware that sharp objects might damage the mesh.

  • It is preferable to use a nylon bristle pan brush with the bristles cut to a length of approximately 25 mm (1″).

  • Brushing should be avoided on sieves finer than 150 microns (No.100), this can cause the wire mesh to loosen.

  • Ultrasonic cleaning is a good way to clean the fine mesh or remove the lodged particles. After this process, ensure that the test sieve is dried thoroughly before use.

We also recommended users to

  • Avoid putting heavy loads onto the sieve surface. Always make sure that the volume of the sample is not going over the maximum permitted volume for each aperture.

  • Organise the stack of sieve with consideration to the steps in aperture sizes , especially when using a sieve shaker. Make sure that there is a larger aperture size sieve nested above the fine mesh sieves. This will help to catch the bigger samples before they land on the finer mesh surface.

  • Average recommended test sieve calibration should be carried out every 12 months. In some cases, every 6 months depending on the level of usage. A weekly check is advised to obtain consistent results. Calibration results will be able to indicate whether the sieve is still accurate to use or it is time to replace with a new one.

If you have chosen to use Glenammer test sieves, congratulations! A satisfied customer recently commented “Glenammer test sieves last three times longer than other brands on the market”. Glenammer offers a life-long warranty against any manufacturing defects.

Glenammer has made the following efforts to produce long-lasting test sieves:

Material: we choose to use high-grade stainless steel to produce our test sieves. Stainless steel is a more durable and stable material for various laboratory testing applications.

Label: we have applied 3D laser technology to generate a permanent metal cut-in label, which will carry the product information on the frame for the whole lifespan.

Reinforced Mesh: For 300 mm diameter (and above) test sieves with apertures 90 micron and under, we have a supporting layer underneath the initial mesh to ensure these fine test sieves last longer.

Posted by Oluwakemi Adi

kemi@aasnig.com, www.aasnig.com

08060874724, 07084594004

HPLC Mobile Phases – 10 bad habits to avoid

1. Measuring the pH of the mobile phase after the organic has been added
pH meters are calibrated to give the correct pH readback in aqueous solution – the buffers you verify this with are aqueous. If you measure the pH with the organic added, the pH will be different to that of measuring before organic addition. However, the most important point is to be consistent.If you do always measure pH after the organic is added, make sure you state this in the method so that everyone does it the same way. It won’t be 100% accurate, but at least it will be consistent. This is probably more important than having the exact pH.

2. Not using a buffer
Buffers are present to control pH and resist a change in pH. Many other parts of method (e.g. sample matrix, CO2 in air, source of water used for your mobile phase) can change the pH of the mobile phase causing shifts in retention, peak shape and peak response. Formic acid, TFA etc. are not buffers.

3. Not using the buffer in its correct pH range
Each buffer salt has a 2 pH unit wide range over which it provides the optimal pH stability. Outside this window the salt is ineffective at resisting change in pH. Either use your buffer within the correct range or pick a buffer whose range covers the pH you require.

4. Adding buffer to organic
Mixing aqueous buffer into the organic phase carries a high risk of the buffer being precipitated – in many cases so finely that it may not be obvious it has happened. ALWAYS add the organic to the aqueous phase, this greatly reduces the risk of buffer precipitation.

5. Using the pump to mix gradients from 0%
Modern pumps are very effective at mixing mobile phases and degassing online, however not everyone who ends up using your method has a high quality pump. Premix your A and B starting mix to a single solution that runs at 100% on line A. e.g. Prepare the starting mixture by mixing 950ml Aqueous with 50ml organic, then filter and degas. This reduces variability between HPLCs, reduces the risk of bubbles and precipitation in the system. Note however that 95:5 mixed on the pump will not give the same retention time as 95:5 premixed in the bottle – you normally need to add a few more percent organic when premixing.

6. Not using the correct pH modifying acid or base for your buffer
Only use the acid or base that forms the buffer salt you are using. E.g. sodium phosphate buffers should be adjusted with only phosphoric acid or sodium hydroxide.

7. Not stating the full information of your buffer in the method e.g. weigh 5g of sodium phosphate into 1000ml of water
The type of buffer (mono, di or tribasic) determines its pH buffering range.
The required molarity is what determines the buffer strength. 5g or anhydrous sodium phosphate and 5g of monohydrate sodium phosphate will have different buffer strengths and will affect retention.

8. Filling lines with organic without checking what was in there before
If the previous method used buffer in line B and your method uses organic in line B there’s a good chance you will precipitate buffer in your pump tubing / pump head and you may cause a lot of damage. If in doubt – flush it out (80:20 water : organic).

9. Propping up bottles to get last drop out
It’s 5 to 5 and you’ve barely got enough mobile phase to finish the run – it’ll be running on fumes by the last few samples. Apart from the risk or running your pump and column dry, mobile phases evaporate from the surface, so the mobile phase at the top of the bottle will have changed composition from the bulk. This portion from the top is exactly what will be running through the column if you use the last dregs in the bottle.

10. Using sonication to degas mobile phase
It’s great for making sure all your buffer salts have dissolved, but it’s the least effective method of degassing AND it quickly heats up the mobile phase causing the organic portion to evaporate. Save yourself problems later – take 5mins to vacuum filter your mobile phase – it degasses and filters in a single step.

Reference

Culled from: LCGC March 08, 2017

Posted by Muyiwa Adebola

muyiwa@aasnig.com, www.aasnig.com

07084594001, 07084594004

The LCGC Blog: Buffers and Eluent Additives for HPLC and HPLC–MS Method Development

Jun 05, 2019

By Tony Taylor

Modern HPLC method development is dominated by a small number of pH adjusting reagents and buffers that are prevalent even when the method uses UV detection. This is driven primarily by the requirements of mass spectrometry. These pH adjusting reagents and buffer combinations are shown in Table I. While they are all, in theory, MS compatible, they are sometimes chosen without justification.

I want to revise some of the basics of HPLC buffer use, as well as highlight some other buffers that can be very useful when these stock buffers aren’t successful. This saves time and money when coming up against roadblocks with separation development as, once all of the usual buffers have been tried, attention turns to changing the column chemistry, which may not be necessary.

Table 1:  Common Eluent pH adjusting reagents and Buffers.

A Revision of Buffers

Remember that the pH adjusting reagents will not provide any buffering capacity, meaning that if changes in pH are encountered by analytes (typically, while the sample diluent and eluent within the instrument tubing or at the head of the HPLC column are mixing) it may result in poor peak shape, poor retention time reproducibility, and potential loss of resolution.

Buffers exhibit their greatest buffering capacity at +/- 1pH unit around the buffer pKa. The closer the eluent pH is to the buffer pKa, the lower the concentration of buffer which needs to be used in order to provide effective resistance to change in pH. For LCMS applications, aim for 10 mM buffer concentration or less and for UV-based applications (less than 25 mM is preferable).

Bear in mind the UV contribution of the additive when working at low UV wavelengths (<220nm) and be smart with UV detector settings to avoid sloping baselines (1,2)

When adjusting the pH of the aqueous portion of the buffer to achieve a pH relative to a known or calculated analyte pKa (for example, when adjusting pH to be well away from the analyte pKa), remember that the pH of the solution will change when the organic component is added. As a very approximate rule of thumb follow these guidelines:

For acidic eluents and buffers

  • +0.22 pH units per 10% acetonitrile
  • +0.15% pH units per 10% methanol

For basic eluents and buffers

  • -0.05 pH units per 10% acetonitrile
  • -0.10 pH units per 10% methanol

Remember that trifluoroacetic acid (TFA) is a strong ion pairing reagent and may severely restrict the detector sensitivity in positive ion mode because the ion pair is strong enough to survive as a neutral complex with the analyte during liberation into the gas phase. Further, TFA is known to linger within mass spectrometer sources and may take prolonged cleaning in order to remove it. TFA may also form strong hydrophobic interactions with silica based HPLC columns, substantively altering the column chemistry. Many users recommend that columns used with TFA are dedicated to separations using this eluent additive.

Ammonium acetate has sparing solubility in acetonitrile and above 60% acetonitrile, vigilance is required to avoid the formation of colorless salt crystals within the eluent reservoir and inner surfaces of the HPLC equipment.

Alternative Buffers and Further Thoughts

The main buffers that can be used as an alternative to TFA are:

  • Pentafluoropropionic acid (PFPA)
  • Heptafluorobutyric acid (HFBA)
  • Methanesulphonic acid (MSA)
  • Ammonium hydrogen carbonate

Given the many unwanted characteristics of TFA, users tend to turn to alternative buffer systems, without realizing that there are several higher perflourinated acids that can be used with MS detection to provide alternative selectivity. They have been widely reported in literature as long ago as 1996 (3,5).

Figure 1: Pentafluoropropionic acid (PFPA, pKa 0.18) and Heptafluorobutyric acid (HFBA pKa 0.4).

Figure 1 shows two such volatile perfluorinated acids that can be used as an alternative to TFA. Retention time under reversed-phase conditions will tend to increase with increasing ion pair chain length; however, care is required to add just enough ion pairing reagent for improved retention. Excess ion-pairing reagent may cause retention loss due to various electrostatic effects associated with adsorption of the ion pairing reagent on the silica surface. These additives are also ion pairing reagents but are weaker than TFA. The ion-pair tends to dissociate within the ESI source, giving rise the corresponding charged analyte in the gas phase. Hence, sensitivity of detection is not affected to the same degree as with TFA. These reagents also linger for much shorter times within ESI sources. Furthermore, each of these reagents will produce an alternative selectivity to the separation carried out with TFA. The acidity of these reagents should also be noted and a stationary phase with good low pH stability should be selected. Column-washing procedures may need to be employed to remove the additives from the stationary phase surface.

We have noted, along with other literature reports (3) that the addition of the perfluorinated acids to the sample diluent can have a marked effect on the peak shape, and sometimes on the retention time stability of the resulting chromatography, especially, when dealing with highly ionogenic compounds.

Methanesulphonic acid (MSA) can also be used as a very effective alternative to TFA when using UV detection. It is a stronger acid than TFA and, as such, will have sufficient capacity at lower concentrations than TFA (a 3-mM solution of MSA gives a similar pH to 0.1% TFA). It will also retain its buffering capacity over a wide-range of acetonitrile concentrations and has the added advantage of a UV cut-off of 195nm. Again, MSA produces altered selectivity to TFA and there are reports that addition of MSA to TFA based eluent systems in HILIC mode can be used to “tune” the selectivity in this separation mode (6)].

Ammonium hydrogen carbonate is an excellent buffer for use at high-pH with good buffering capacity over pH 8-11 and possibly wider at higher ionic strength. The extended buffering range is due to the ammonia—ammonium buffering capacity being additive to the hydrogen carbonate-carbonate capacity in what is traditionally called a “mixed buffer.”  That is, the pKa of the two buffering systems are relatively close. Compare this to the use of ammonium acetate or formate buffers at low pH where the buffering ranges of the ammonium species and the format or acetate are several pH units apart (see Table I). Under these circumstances, the ammonium ion is merely acting as an MS-friendly counter ion in place of sodium or phosphorous ions. Ammonium hydrogen carbonate is MS-friendly and has a UV cut-off of 190 nm.

These buffers can produce somewhat unstable retention at pH 7; however, this is thought to be due to the less effective buffering capacity in the “valley” region between the first and second pKa values of the buffer. Eluents above pH 8 should produce very effective buffering.

Thinking of separations at high pH brings us to another interesting point when selecting buffers for the separation of basic analytes with MS detection. One might expect that selecting an eluent pH in which the analyte is expected to be in the neutral form (eluent pH above the analyte pKa for basic analytes) will lead to reduced analyte detector response. However, this has been shown not to be the case in many examples and has come to be known as “wrong way round” ionization (7,8). This mode of ionization is reported to be driven by process such as:

  • Low ionic strength solutions at near neutral pH—electrolytically-produced protons are abundant at the droplet surface from which analyte ions are desorbed
  • In strongly basic ammonia solutions, gas phase protons transferred from ammonium ions can be the dominant charging mechanism
  • Discharge induced ionization also occurs with high ionic strength at neutral or high pH

Therefore, It is incumbent upon us to explore separations involving basic analytes at high pH to gain alternative selectivity, even if this appears to be counterintuitive to theory.

One further note on MS signal intensity is the use of forced adduct formation to improve the sensitivity of the analyte, or to distinguish one analyte from another within the MS chromatogram. Solutions of sodium or ammonium acetate (for example) can be infused into the eluent flow post-column in order to promote adduct formation, which is often attendant with an increase in analyte signal. Typically, 1–5-mM solutions are used to prevent source contamination or blockage and only the purest reagents available should be used.

Finally, one very useful eluent additive was recently reported (9), helping to overcome the effects of analyte binding to the metal surfaces within the HPLC system as well as improving the peak shape and detector sensitivity for anionic analytes. Medronic acid (Figure 2) can be used as a very useful alternative to EDTA with LC–MS analysis and has been shown to produce much lower degrees of ion suppression.

Figure 2: Medronic (Methylenediphosphonic) acid (pKa – 1.27)

This is especially useful in the analysis of peptides and proteins, and typically 5 mM of medronic acid can be added to buffered mobile phase (ammonium acetate, for example) to provide highly-effective “deactivation” resulting in improved peak shapes, detector sensitivity, and quantitative reproducibility.

I would urge anyone developing an HPLC method to consider whether they are selecting the most appropriate eluent additive prior to commencing laboratory work and to be open to exploring buffers beyond the established norm.

On the frontline of the selectivity battle, we need to have as many weapons as possible!

References

  1. So just how well set-up is your UV detector?
  2. UV detection for HPLC – Fundamental Principles, Practical Implications
  3. Flieger, J Chromatogr A, 1217, 540–549 (2010).
  4. Shibue, C.T. Mant, R.S. Hodges, J Chromatogr A,, 1080, 68–75 (2005).
  5. D. Pearson*, Mark C. McCroskey, J Chromatogr A, 746, 277–281(1996).
  6. V. McCalley, J Chromatogr A, 1483, 71–79 (2017).
  7. A. Mansoori, D.A. Volmer, R.K. Boyd, Rapid Commun. Mass Sp. 11 1120–1130 (1997).
  8. Zhou, S., Cook, K.D.: Am. Soc. Mass Spectrom. 11, 961–966

Written by Akinbuli Opeyemi,

www.aasnig.com, opeyemi@aasnig.com

07084594004, 08068129603

Gas management system for Gas chromatography laboratory and importance of changing filters regularly.

A clean gas stream is critical to the quality of your GC analysis and the reliability of your analytical results. Carrier gas must contain less than 1 ppm of oxygen, water vapor, and other trace contaminants for stable baselines on all detectors and to prevent column degradation, shortened column lifetime, and increased stationary phase bleed. Clean fuel gases and make-up gases are also essential for stable detector baselines. Your gas management system must deliver a high-purity gas stream from your source to your instrumentation without introducing contaminants. Figure below shows a typical gas management system using a cylinder

Questions to consider when building gas management system for a GC Lab:

  1. What gases do you require and at what purity?
  2. What flow rates and/or pressures are required?
  3. Are you using gas generators or high-pressure cylinders?
  4. What type of tubing and fittings are needed to build your system?
  5. What type of gas purifiers should you install in your system?

This article explains why it is important to replace your gas filters annually instead of waiting for the indicators to change color

A significant number of instrument and column complaints are carrier gas related, which could be caused by breakthrough from filters that are not replaced in time.

The possible consequences of a filter breakthrough are:

  • Gas distribution system behind the filter will be contaminated (fast cleaning nearly impossible, bleeds for months)
  • Instrument gets contaminated, expensive maintenance required
  • Column lifetime reduced, bad analytical results, high cost of ownership, unnecessary changing of column brands
  • MS source gets contaminated, expensive maintenance required, long system shutdown.

Figure showing reduced instrument baseline due to filters

Gas Purifiers

Gas purification is essential in your gas management system. Carrier gases must contain less than 1 ppm of oxygen, water vapor, or any other trace contaminant to prevent column degradation, shortened column lifetime, and increased stationary phase bleed. Contaminants cause ghost peaks to appear during temperature programming and degrade the quality of analytical data. The expense of using high-purity gases in combination with carrier gas purifiers will be offset by longer column lifetime and less instrument maintenance along with better instrument sensitivity. Gas purifiers are available for specific types of contamination (moisture, hydrocarbon, or oxygen) or as a combination of filters that provides broader protection. These purifiers can be installed in-line or using a quick-install baseplate system.

A typical GC and GC-MS laboratory gas filter system contains three types of filtering media:

  • Oxygen Catalyst for absorbing traces of oxygen
  • Activated Carbon for adsorbing hydrocarbons
  • Molecular Sieve for adsorbing moisture

These filter media types can be divided into two groups:

  • Chemically absorbent media:

Like a Venus fly trap, when it comes in contact with a contaminate it absorbs it and does not let it go. Adsorption differs from absorption, which also removes things, but the result is swelling of the media. The media size increase equals the amount of material removed

  • Surface adsorbent media

Like a floor mop, where the contaminant is trapped onto the surface of the media and retained. The material doing the adsorption does not change in size.

Also note that most filters do not provide an indicator for hydrocarbons, so you would never know whether the activated carbon has reached its adsorption capacity.

Filter Media Breakthrough

Both adsorption and absorption media have fixed capacities, meaning they hold just so much, since they are storing the material removed from the gas, not destroying it.

Normal situation

Hydrocarbon breakthrough

Over time, the pores of the activated carbon fill up. The molecules that are adsorbed with higher energy (larger mass) can displace the lower-energy molecules that are less tightly held. This phenomenon, called displacement, may knock the smaller particles off the media, straight to the column and into the instrument.

Moisture breakthrough

The molecular sieve adsorbs moisture until it cannot adsorb anymore. If the humidity level of the gas is lower than in the molecular sieve, it will de-adsorb its moisture until it is in “balance” with the lower humidity level of the gas, which means the filter could increase the amount of moisture in your gas.

Oxygen breakthrough

The media size increase equals the amount of material removed. When the media has reached its absorption capacity it will not release the already trapped contaminants but it will also not absorb any new ones.

When am required to change my gas filter?

Filter Media Breakthrough Indicators

The visual indicators are mainly for urgent situations such as a leak or high amount of impurities breaking through.

Indicators are typically placed behind the filter media bed. When they change color shortly after installation of the filter it usually means there is a leak or that the gas distribution system including the manifold to which the filter is connected was not flushed properly prior to – or during – installation of the filter.

When the indicator changes color during normal operation of the filter it indicates that the filtration media has reached its capacity, and the filter should be replaced immediately to avoid contaminant breakthrough.

Other components such as branched hydrocarbons which are also trapped by the molecular sieve are not shown by the moisture indicator.

Also note that most filters do not provide an indicator for hydrocarbons, so you would never know whether the activated carbon has reached its adsorption capacity.

 

Conclusion

Replace your filter before the indicators start changing color to prevent breakthrough and to avoid high maintenance and repair cost of your instrument.

As explained previously, filters which are not maintained on a regular interval can cause the outgoing gas to become more contaminated than the original source gas. The color indicators used in gas filters are so called ‘last minute’ indicators and require quick action. You can compare it to the engine oil indicator in your car. When the engine oil indicator is blinking on your dashboard, the car should not be driven and ignition should be switched off unless topped with engine oil – In event of taking risk to drive – high probability of engine getting seized causing high expenditure to repair or replace the engine.

The same is valid for gas filters. When one of the indicators starts to change color, the filter should not be used and instrument should be switched off unless the filter is immediately being replaced with a new one – In event of taking risk to continue – there will be a high probability of contaminant breakthrough causing high expenditure on instrument maintenance as compared to planned annual preventive maintenance on your instrument and gas filters.

So what you can do as an end-user is to always buy an additional set of filters to keep on standby in case the indicator starts changing color or to use a good preventive maintenance plan or tool (such as the electronic indicator) to replace your filters at least once a year.

Reference

https://www.peakscientific.com/articles/use-of-traps-in-gc/

https://www.restek.com/Technical-Resources/Technical-Library/General-Interest/general_GNSS1758B-UNV

https://www.shimadzu.fr/sites/shimadzu.seg/files/shimadzu_brochure_filtre_gaz_super-clean_c180e083a_-_2017-06.pdf

Written by Oweh Gabriel,

www.aasnig.com, gabriel@aasnig.com

08035696303

Sustainability in the Workplace: How to Reduce the Use of Plastic

The attention placed on the plastic problem has seen a reduction in the use of plastic straws, retailers ending the sale of single-use bottles of carbonated drinks, and other eco-friendly measures to kerb the amount of plastic we use and throw away. Progress made in the reduction of plastic has been slow, and more must be done to reduce the devasting impact plastic can have on our environment.

use of plastic in the workplace

Wildlife is particularly vulnerable to plastic and can easily become entangled in discarded items, or get a piece lodged in their digestive tracts, leading to injury or death. Certain plastics also release harmful chemicals into the surrounding soil, while the breakdown of biodegradable plastics can release methane, a greenhouse gas that’s one of the key triggers of global warming.

Recent reports on recycling have shown increases in the amount of plastic being repurposed year on year, especially compared to other materials like glass, textiles, paper and card. But, while this does signify a move towards more environmentally-conscious practices, figures suggest that the increase in the amount of plastic being recycled is down to a larger amount of plastic containers in the first place. Because of this, local authorities have added plastic bottles to their range of materials alongside paper, card, glass and metals, while a good majority have also started collecting pots, tubs and trays as of 2016/17.

As a result of more plastic packaging and an increased awareness for the importance of recycling, Welsh councils have nearly doubled the amount of plastic they collect per person, while Northern Irish authorities have seen an increase of more than a third. All told, the increase in plastic recycling can be seen in the Eunomia’s Carbon Index, which gives an idea of the local councils delivering the greatest carbon benefits. Currently, Cheshire West and rank the highest, with Lewisham being the worst performing local authority for the amount of plastic recycled per household.

Reference

www.mynewlab.com

Written by Oluwakemi Adi

08060874724, 07084594004

kemi@aasnig.com, www.aasnig.com

 

MAINTAINING A FLAME IONIZATION DETECTOR: WHEN AND HOW TO CLEAN

Noisy chromatograms, random spikes, and poor detector sensitivity are symptoms of a dirty FID — a common problem in gas chromatography. You will consistently obtain better chromatograms and reduce instrument downtime if you keep the FID clean.

The most common source of contamination in a flame ionization detector (FID) is bleed from silicone stationary phases and silylating reagents, which combust in the FID and produce silica. When deposited on surfaces within the detector, this white powder causes noisy chromatograms, random spikes, and poor detector sensitivity Figure below;

Figure 1. A Noisy Chromatogram, Caused by a Dirty FID

In this article we will

  1. methods of troubleshooting noise to confirm whether the source is a dirty FID,
  2. method of cleaning FIDs, and
  3. ways to reduce contamination in the future
    IS THE DETECTOR REALLY THE PROBLEM?
    Before you shut down your instrument and clean your detector, it is wise to confirm that the problem is detector-related, rather than related to some other component of your system. The few, simple procedures described here can eliminate other possibilities as the source of the problem.
  • Carrier Gas and Stationary Phase

Seal the detector inlet in the oven with a  plug and ignite the detector. If the chromatogram noise disappears, then the source of the problem is contaminants in the carrier gas or bleed from the chromatography column, not a dirty FID.

  • Hydrogen and Air Systems

Hydrogen and air used in the sulfide can be a source of contamination, especially when problems emerged after replacing the cylinder. An incorrect flow rate in either source can cause noise, lack of sensitivity, and/or difficulty when igniting the flame. A contaminated cylinder of gas could be the source of the problem, especially if the noise appeared several hours after you changed a cylinder. Check each cylinder for contaminants and replace if necessary. To eliminate the problem of contaminated air, we recommend using a zero air generator

  • Electrical System

Electrical interference may exhibit similar symptoms dirty FID. There may be a defect electrometer, poor contact or interference by other devices in the lab. To isolate this source of noise, disconnect the electrometer cable(s) from the FID. If noise persists, it is coming from the electrical system.

Precautions before cleaning

  • Isolate electrical source, make sure to unplug the power cord!
  • Remember that the detector may be hot! be sure the collector assembly is cool before you begin
  • When dismantling FIDU pay attention insulating parts. Use tweezers, whether these parts do not transfer dirt from your hands or gloves. Beware of possible scratches.
  • Pay Attention ! Carefully note the distance from the collector assembly to the flame jet.

How to Clean an FID
To properly clean an FID, you must clean the
1. Collector assembly
2. The jets:
Jet cleaning  procedure.

  • Run a cleaning wire through the top of the jet. Run it back and forth a few times until it moves smoothly. Be careful not to scratch the jet. (Do not force too large a wire or probe into the jet opening or the opening will become distorted. A loss of sensitivity, poor peak shape and/or lighting difficulties may result if the opening is deformed.)
  • Fill an ultrasonic cleaning bath with aqueous detergent, and place the jet in the bath. Sonicate for five minutes.
  • Use a jet reamer to clean the inside of the jet.
  • Sonicate again for five minutes.
    NOTE: From this point on, handle the parts only with forceps!
  • Remove the jet from the bath and rinse it thoroughly, first with hot tap water and then with a small amount of GC-grade methanol.
  • Blow the jet dry with a burst of the compressed air or nitrogen, and then place the jet on a paper towel and allow it to air dry.
  1. The Teflon® or ceramic insulators: Ceramic parts of an FID are best cleaned with aqua regia, a 1:~3 mixture of concentrated nitric and hydrochloric acids, at ambient or mildly elevated temperature. Before treatment, remove all metal and rubber from the ceramic parts – aqua regia will attack these.
  2. The housing.

After you have cleaned all parts of the detector, check all Orings and replace them if necessary. Worn-out O-rings will cause gas leaks, which can produce detector noise or an increase in detector contamination. Reassemble the FID, light the flame, and allow the detector temperature to equilibrate at 10°C–50°C higher than the column will reach during typical operation. This will reduce the amount of phase condensing onto the detector parts. Do not exceed the maximum temperature limit of the stationary phase – many columns fit far enough into the detector to expose the phase to these elevated temperatures. Set the proper flow rates for hydrogen and compressed air (refer to the instrument manual), and ignite the flame. Turn on the electrometer and allow a few minutes for warmup. The flame should now be stable and noise-free.

REDUCING DETECTOR NOISE AND CONTAMINATION

  1. Conditioning:
    Most detector noise and contamination is the result of column bleed. The amount of bleed is greatest when the column is initially conditioned. Your detector will remain clean longer if you condition a new column before connecting it to the detector. Byproducts eluted during conditioning, potentially harmful to the FID, are voided into the oven. Connect the column inlet to the injector as usual. Place a restrictor at the column exit to prevent back diffusion of air into the column (exposure of a heated column to air can destroy the liquid phase). Purge the column with carrier gas at room temperature for a few hours before you begin the temperature program. Do not allow a combustible carrier gas such as hydrogen, methane, etc. to exit the column into the oven. Pipe these materials out of the oven and into a hood. (Be sure to attach a restrictor to the outlet of the pipeline in the hood.) Consult the column manufacturer for conditioning details, i.e., duration and temperature of conditioning. Do not routinely condition new columns at the maximum temperature limit of the stationary phase – this will reduce column life. Connecting a well-conditioned column to a clean FID should produce good sensitivity. If detector stability quickly degenerates, you should evaluate the quality of your stationary phase and carrier gas.
  2. Stationary Phase
    Use GC quality stationary phases whenever possible — they are purified to remove lower molecular weight components. Technical grade materials will bleed more than GC quality materials.
  3. Carrier Gas
    Moisture and oxygen in the carrier gas will cause stationary phase to deteriorate and bleed. Use chromatography-quality gases, and periodically monitor the gas system for leaks, which might allow atmospheric oxygen and water to enter the column.
  4. Septa
    Frequently check the septum for leaks. A leaking septum can allow oxygen and water to enter the carrier gas and cause the stationary phase to deteriorate and bleed. To check for leaks without contaminating the septum (and subsequent samples) with liquid leak detectors, use an electronic GOW-MAC leak detector, If you run your instrument frequently, we recommend you change the septum regularly.

    Reference
    https://www.agilent.com/en/support/gas-chromatography/kb001441https://www.chromservis.eu/i/32606/g/hints-and-tips
    https://www.sigmaaldrich.com/content/dam/sigma-aldrich/docs/Supelco/Bulletin/4488.pdf
    http://hiq.linde-gas.com/en/analytical_methods/gas_chromatography/flame_ionisation_detector.html

Written by Oweh Gabriel,

www.aasnig.com, gabriel@aasnig.com

08035696303

 

 

 

 

 

Know Your Sample: Size Matters

BY: Douglas E. Raynie

Over the past several years, there has been a trend toward preparing increasingly smaller samples. In many cases, this approach was taken to demonstrate that extractions and sample handling procedures at the microscale and smaller is possible. With the current emphasis on bioanalytical and related technologies, even greater legitimacy is given to these approaches. Hence, the advent of dried-blood spot (DBS) analyses and other approaches. One approach to microsampling for bioanalyis is solid-phase microextraction (SPME), also referred to as bio-SPME, which is discussed here.

Sampling and Sample Heterogeneity

My thoughts as I heard of recent, somewhat controversial, developments in finger-prick sampling for blood tests were concern over the statistics of sample size and homogeneity. Most analysts are widely aware that the standard deviation of sampling and analysis increases with decreasing analyte concentration. Horwitz (1) evaluated interlaboratory validation studies and developed the “trumpet” shown in Figure 1. Although some bias is evident in every sampling protocol, when Meyer (2) presented the relationship between sampling and measurement uncertainty in 2002, she claimed that deviations from the Horwitz curve were caused by the sample matrix and the sample preparation procedure. Meyer provided the following advice: Avoid all possible sources of contamination with trace analysis; use large volumes when possible since smaller volumes are difficult to handle and loss of sample material is less severe; mass-based measurements are often more reproducible than volumetric measurements; and use minimal sample handling steps with small-volume samples. Others have also demonstrated the relationship between sampling precision and sampling size. For example, Thiex and colleagues (3) reported the expected relative standard deviation from laboratory subsampling as a function of maximum particle size within the sample. As presented in Table I and substantiated by the Horwitz relationship, one cannot simultaneously have good sampling precision and small samples.

Figure 1: The Horwitz “trumpet” displaying the inverse relationship between analyte concentration and relative standard deviation of sampling. (Adapted from reference 2.)

Moving beyond sample homogeneity concerns, the Royal Society of Chemistry’s Analytical Methods Committee explored representative sampling from an analytical and statistical viewpoint (4). They prefer the term “appropriate sampling” to “representative sampling.” They made this distinction because of the survey statistics definition of representative sampleas “a sample for which the observed values have the same distribution as that in the population,” while the analytical definition states “a sample resulting from a sampling plan that can be expected to reflect adequately the properties of interest in the parent population.” This concept of adequacy in the analytical definition implies an inherent sampling bias and recognizes that in many, especially regulatory, cases analytical results are compared with a limit value. This limit value is often a “fitness for purpose,” which allows the use of analytical results to be used in decision making. Appropriateness of sampling can be improved by increasing the sample size or the number of samples.

Note that these relationships between sample size and heterogeneity are primarily derived from investigations of solid samples, including food and feeds. However, most microsampling applications are used in bioanalysis, especially those involving blood samples. For reasons of diffusion and turbulent flow, liquid samples can be assumed to be considerably more homogeneous than solid samples.

Overview of Microsampling in Bioanalysis, Including Bio-SPME

Along with analysis of DBS, paper-based and more-conventional microfluidic approaches are gaining popularity. Such procedures are simple, inexpensive, and easy to use. A balance of hydrophobic and hydrophilic treatments controls fluid movement in these devices, resulting in their claimed reliability. One significant advantage of these approaches is their applicability outside of the laboratory, including nonclinical settings, though sample drying of blood spots can present a concern. Capillary microsampling allows collection of microliter sample volumes along with subsequent steps such as separating plasma and serum. These approaches will be the subject of a future “Sample Prep Perspectives” column.

Another sample preparation trend we’ve noticed is interest in SPME, especially since the lapse of patent protection of the initial products. In the case of conventional SPME, a stationary phase, usually a gas chromatography (GC)-type phase, is coated onto a fused-silica fiber encased in a syringe-needle device. The coated fiber is exposed to the sample by either immersion in a liquid sample or exposure to a vapor sample. The adsorbed sample is then desorbed either thermally in a GC inlet or via solvent rinsing into a liquid chromatograph.

Biocompatible SPME (bio-SPME) is a microsampling approach for bioanalysis based on SPME, but featuring some key differences. With bio-SPME, functionalized silica particles are embedded in an inert binder that is coated or bonded onto metal fibers. The use of the binder minimizes interferences from biomacromolecules. Bio-SPME is available in hypodermic needle and pipette tip formats. Like conventional SPME, the approach is not exhaustive and relies on an equilibrium between the analyte in the biofluid and the fiber materials. Figure 2 displays the kinetics of bio-SPME sampling, which are similar to conventional SPME. Initially, a rapid adsorption of the analyte onto the functionalized silica is observed, followed by an asymptotic approach to the equilibrium amount of analyte isolated.

Figure 2: The kinetics of the bio-SPME process, demonstrating an asymptotic approach to quantitative equilibrium. (Courtesy of Supelco.)

Two particular advantages of bio-SPME are of special interest. First, the device can be directly inserted into small animals for sampling at or near the point of interaction during physiological studies. This allows multiple analyses per animal, since the animal is not sacrificed, leading to more cost-effective studies and more reliable results since there are multiple analyses per animal. Relative standard deviations around 30% demonstrate the need to strongly consider the uncertainty considerations presented by Horwitz and discussed earlier. The second major advantage is the direct ionization of analytes on the bio-SPME fibers for mass spectrometry, as demonstrated in Figure 3 (5). This schematic shows the ionization occurring when the fiber and spray tip are sharp and a spray solvent carries the analyte into a high-voltage region to create an electric field between the bio-SPME device and the inlet to the mass spectrometer. Quantitative results are similar to other reports of bio-SPME and are 5–10x better than with DBS analysis. Spray solvent flow rates, positioning of the fiber, and other parameters are being optimized.

Conclusions

Microsampling for bioanalysis and other applications is gaining in popularity. One new technique in this area is the reapplication of the SPME approach, designed for biological applications. However, in all microsampling approaches, measurement uncertainty and sample homogeneity concerns must always be considered.

References

  1. W. Horwitz, L.R. Kamps, and K.W. Boyer, J. Assoc. Off. Anal. Chem. 63,1344–1354 (1980).
  2. V. Meyer, LCGC North Am. 20(2), 106–112 (2002).
  3. N. Thiex, L. Novotny, C. Ramsey, G. Latimer, L. Torma, and R. Beine, Guidelines for Preparing Laboratory Samples (Association of American Feed Association Officials, 2008).
  4. M.H. Ramsey and B. Barnes, Anal. Methods 8, 4783–4784 (2016).
  5. S. Ahmad, M. Tucker, N. Spooner, D. Murnane, and U. Gerhard, Anal. Chem87, 754–759 (2015).

Written by Akinbuli Opeyemi,

www.aasnig.com, opeyemi@aasnig.com

07084594004, 08068129603

Lab Safety Rules and Guidelines

Lab safety rules

Having a strong set of overall laboratory safety rules is essential to avoiding disasters in the lab. Laboratories recently scoured the safety policies of several laboratories to determine some of the most common lab safety rules out there, to help you whether you’re developing or updating a set of policies for your own lab. Of course, safety rules are only effective when they are enforced, which is why strong lab management is so important to a safe laboratory as well. Knowing the proper laboratory safety signs and symbols is also important.

Here are the safety rules that most commonly came up in our look at several laboratories’ policies:

General lab safety rules

The following are rules that relate to almost every laboratory and should be included in most safety policies. They cover what you should know in the event of an emergency, proper signage, safety equipment, safely using laboratory equipment, and basic common-sense rules.

  1. Be sure to read all fire alarm and safety signs and follow the instructions in the event of an accident or emergency.
  2. Ensure you are fully aware of your facility’s/building’s evacuation procedures
  3. Make sure you know where your lab’s safety equipment—including first aid kit(s), fire extinguishers, eye wash stations, and safety showers—is located and how to properly use it.
  4. Know emergency phone numbers to use to call for help in case of an emergency.
  5. Lab areas containing carcinogens, radioisotopes, biohazards, and lasers should be properly marked with the appropriate warning signs.
  6. Open flames should never be used in the laboratory unless you have permission from a qualified supervisor.
  7. Make sure you are aware of where your lab’s exits and fire alarms are located.
  8. An area of 36″ diameter must be kept clear at all times around all fire sprinkler heads.
  9. If there is a fire drill, be sure to turn off all electrical equipment and close all containers.
  10. Always work in properly-ventilated areas.
  11. Do not chew gum, drink, or eat while working in the lab.
  12. Laboratory glassware should never be utilized as food or beverage containers.
  13. Each time you use glassware, be sure to check it for chips and cracks. Notify your lab supervisor of any damaged glassware so it can be properly disposed of.
  14. Never use lab equipment that you are not approved or trained by your supervisor to operate.
  15. If an instrument or piece of equipment fails during use, or isn’t operating properly, report the issue to a technician right away. Never try to repair an equipment problem on your own.
  16. If you are the last person to leave the lab, make sure to lock all the doors and turn off all ignition sources.
  17. Do not work alone in the lab.
  18. Never leave an ongoing experiment unattended.
  19. Never lift any glassware, solutions, or other types of apparatus above eye level.
  20. Never smell or taste chemicals.
  21. Do not pipette by mouth.
  22. Make sure you always follow the proper procedures for disposing lab waste.
  23. Report all injuries, accidents, and broken equipment or glass right away, even if the incident seems small or unimportant.
  24. If you have been injured, yell out immediately and as loud as you can to ensure you get help.
  25. In the event of a chemical splashing into your eye(s) or on your skin, immediately flush the affected area(s) with running water for at least 20 minutes.
  26. If you notice any unsafe conditions in the lab, let your supervisor know as soon as possible.

Housekeeping safety rules

Housekeeping lab safety rules

Laboratory housekeeping rules also apply to most facilities and deal with the basic upkeep, tidiness, and maintenance of a safe laboratory.

  1. Always keep your work area(s) tidy and clean.
  2. Make sure that all eye wash stations, emergency showers, fire extinguishers, and exits are always unobstructed and accessible.
  3. Only materials you require for your work should be kept in your work area. Everything else should be stored safely out of the way.
  4. Only lightweight items should be stored on top of cabinets; heavier items should always be kept at the bottom.
  5. Solids should always be kept out of the laboratory sink.
  6. Any equipment that requires air flow or ventilation to prevent overheating should always be kept clear.

Dress code safety rules

Dresscode lab safety rules

As you’d expect, laboratory dress codes set a clear policy for the clothing employees should avoid wearing in order to prevent accidents or injuries in the lab. For example skirts and shorts might be nice for enjoying the warm weather outside, but quickly become a liability in the lab where skin can be exposed to heat or dangerous chemicals.

  1. Always tie back hair that is chin-length or longer.
  2. Make sure that loose clothing or dangling jewelry is secured, or avoid wearing it in the first place.
  3. Never wear sandals or other open-toed shoes in the lab. Footwear should always cover the foot completely.
  4. Never wear shorts or skirts in the lab.
  5. When working with Bunsen burners, lighted splints, matches, etc., acrylic nails are not allowed.

Personal protection safety rules

Personal protection lab safety rules

Unlike laboratory dress code policies, rules for personal protection cover what employees should be wearing in the lab in order to protect themselves from various hazards, as well as basic hygiene rules to follow to avoid any sort of contamination.

  1. When working with equipment, hazardous materials, glassware, heat, and/or chemicals, always wear face shields or safety glasses.
  2. When handling any toxic or hazardous agent, always wear the appropriate gloves.
  3. When performing laboratory experiments, you should always wear a smock or lab coat.
  4. Before leaving the lab or eating, always wash your hands.
  5. After performing an experiment, you should always wash your hands with soap and water.
  6. When using lab equipment and chemicals, be sure to keep your hands away from your body, mouth, eyes, and face.

Chemical safety rules

Chemical lab safety rules

Since almost every lab uses chemicals of some sort, chemical safety rules are a must. Following these policies helps employees avoid spills and other accidents, as well as damage to the environment outside of the lab. These rules also set a clear procedure for employees to follow in the event that a spill does occur, in order to ensure it is cleaned up properly and injuries are avoided

  1. Every chemical should be treated as though it were dangerous.
  2. Do not allow any solvent to come into contact with your skin.
  3. All chemicals should always be clearly labeled with the name of the substance, its concentration, the date it was received, and the name of the person responsible for it.
  4. Before removing any of the contents from a chemical bottle, read the label twice.
  5. Never take more chemicals from a bottle than you need for your work.
  6. Do not put unused chemicals back into their original container.
  7. Chemicals or other materials should never be taken out of the laboratory.
  8. Chemicals should never be mixed in sink drains.
  9. Flammable and volatile chemicals should only be used in a fume hood.
  10. If a chemical spill occurs, clean it up right away.
  11. Ensure that all chemical waste is disposed of properly.

Chemistry lab safety rules

As chemistry labs are one of the most common types, these basic chemistry lab safety rules are relevant to many scientists, dealing with the safe performance of common activities and tasks in the average chemistry lab:

  1. Before you start an experiment, make sure you are fully aware of the hazards of the materials you’ll be using.
  2. When refluxing, distilling, or transferring volatile liquids, always exercise extreme caution.
  3. Always pour chemicals from large containers to smaller ones.
  4. Never pour chemicals that have been used back into the stock container.
  5. Never tap flasks that are under vacuum.
  6. Chemicals should never be mixed, measured, or heated in front of your face.
  7. Water should not be poured into concentrated acid. Instead, pour acid slowly into water while stirring constantly. In many cases, mixing acid with water is exothermic.

Electrical safety rules

Electrical lab safety rules

Like almost every other workplace, laboratories contain electronic equipment. Electrical safety rules help prevent the misuse of electronic instruments, electric shocks and other injuries, and ensure that any damaged equipment, cords, or plugs are reported to the appropriate authorities so they can be repaired or replaced.

  1. Before using any high voltage equipment (voltages above 50Vrms ac and 50V dc), make sure you get permission from your lab supervisor.
  2. High voltage equipment should never be changed or modified in any way.
  3. Always turn off a high voltage power supply when you are attaching it.
  4. Use only one hand if you need to adjust any high voltage equipment.  It’s safest to place your other hand either behind your back or in a pocket.
  5. Make sure all electrical panels are unobstructed and easily accessible.
  6. Whenever you can, avoid using extension cords.

Laser safety rules

Laser lab safety rules

Perhaps not as common as some of the other laboratory safety rules listed here, many laboratories do use lasers and it’s important to follow some key rules of thumb to prevent injuries. In particular, accidents due to reflection are something that many employees may not think about. A clear set of rules for the use of lasers is essential to ensure that everyone is aware of all hazards and that the appropriate personal protective equipment is worn at all times.

  1. Even if you are certain that a laser beam is “eye” safe or low power, you should never look into it.
  2. Always wear the appropriate goggles in areas of the lab where lasers are present. The most common laser injuries are those caused by scattered laser light reflecting either off the shiny surface of optical tables, the sides of mirrors, or off of mountings. Goggles will help you avoid damage from such scattered light.
  3. You should never keep your head at the same level as the laser beam.
  4. Always keep the laser beam at or below chest level.
  5. Laser beams should never be allowed to spread into the lab. Beam stops should always be used to intercept laser beams.
  6. Do not walk through laser beams.

References

www.lab-health-and-safety

written by Oluwakemi Adi

kemi@aasnig.com

08060874724.

Maximize Performance and Minimize Maintenance with Genuine Agilent LC Spares

Introduction
Regular replacement of crucial parts can help keep your HPLC systems at optimum performance, reduce system downtime and repair costs, as well as extend your instrument’s life time. However, these benefits can only be achieved when using high-quality parts that are durable, clean, and fit perfectly into the system. We inspected different LC instrument spare parts from Agilent and other vendors.
The results show deficiencies in parts from other vendors, including:
• Inconsistent materials
• Contamination issues
• Shorter life time
• Outdated design
Therefore, use of parts not from Agilent could cause premature instrument failure, increase downtime, and deliver inaccurate or false results.

Solvent inlet filters
Solvent inlet filters represent the first barrier for retaining particulates, precipitation, microbes from mobile phases,
buffers, and salt solutions. Filters are significant in preventing system blockage, pressure increase, and contamination.

Cleanliness
Cleanliness of parts is vital for avoiding system contamination. Agilent solvent filters are packed in ultraclean antistatic bags with an inner metallic coating that does not release contaminants such as plasticizers or antioxidants.
LC/MS analysis shows that filters not from Agilent, packed in normal plastic packs, can cause extra peaks during analysis. Erucamide, a common slip agent used in polyethylene films, is one such example.

Pore size
A good solvent glass filter should have a defined, homogenous pore size to effectively block particulates above a certain size, while letting mobile phases through without significant pressure increase. Too large pore size leads to deficiency of filtration, while pores that are too small can cause pressure increase, resulting in solvent pumping difficulties. Inspection of Agilent and other vendor solvent glass filters by scanning electron microscopy (SEM) shows uniform pore sizes and smooth particle surfaces in the Agilent filter. In contrast, other vendor filters had inconsistent particle and pore sizes. The small particles or particle fragments shown on the other vendor filter could be flushed into the flow path, blocking the pump frit, capillaries, valves,
or columns.

PTFE frits
The PTFE frit is another crucial part in the flow path that prevents particulates and microbes from getting into the system. It is important that frits maintain their shape up to the pressure limit of the system, since collapse or abrasion of the frit can release PTFE particles, resulting in blockage or loss of analysis efficiency. SEM inspection reveals. If the frit is abraded,particles that are too large can block the flow path, while
particles too small can pass through the column inlet frit, getting into the column, or even reach the detector causing
contamination of the flow cell. In contrast to alternative frits, Agilent frits are designed to have a defined particle size to avoid these issues.

Pump piston seals
The piston seal is an essential part of the pump that directly impacts its performance, which depends on many design
characteristics.

Spring tightness
Seal springs have to apply a constant force that complies with the tolerances of the instrument. Springs that are too soft can cause diffusion of air bubbles into the pump head, resulting in pressure ripple and air in the column. Springs too hard can lead to more abrasion between seal and piston, leading to significant decrease in the seal’s lifetime. Agilent seals use specially designed springs that maintain optimal strength to ensure perfect sealability and longevity. Comparison with third-party seals clearly shows differences in size, space, and density of the spring coils.

Material
Agilent seals are manufactured from a proprietary polymer blend with optimized elasticity, firmness, and hydrophobicity, which have large effects on pressure ripple, cold-flow behavior of solvents, and removal of air bubbles. Agilent seals also feature optimal functionality at a wide temperature range from 4 to 60 °C, to adapt to different temperature conditions in different regions. Another feature is that we use specific copper-free manufacturing tools instead of common brass tools, to avoid copper contamination of the system. Therefore, use of third-party seals that clearly have different designs, materials, and features than Agilent seals can result in high risk of compromising your instrumental and analytical efficiency. Agilent uses an ultraclean plastic cap to protect the needle tip from collision and abrasion, contamination, and blockage through particulates. In comparison, there was no proper protection on the third-party needle.

Outlet check valves
The outlet check valve has profound impact on pressure stability and pump flow. The valve has to work quickly,
accurately, and reliably to achieve a precision eluent flow without disturbance such as pressure drop or pressure ripple.The original design of the Agilent outlet check valve had a cylindrical seat and separate gold seal. The cylindrical seat was limited in its resistance to high pressure and alternating pressure loads, resulting in limited life time. In addition, the separate gold seal cap could be deformed by pressure loads due to the ductility of gold, causing leakage so that cap had to be retightened and sometimes changed.
Agilent therefore developed a new design for its outlet check valve to enhance durability and reliability. The new generation outlet valve has a unique double-coned seat to resist the highest pressure ranges, as well as an integrated gold‑plated seal to minimize tolerance of seal edge geometry. In addition, since a gold seal cap is no longer required, there is no need to change the gold seal, which makes this part maintenance free. In comparison, the outlet check valve from another vendor still uses the older design, resulting in risks of higher pressure ripple, poorer flow and retention time precision, as well as shorter life time and more maintenance.

Injection needles and needle seats.

The injection needle and needle seat need to match perfectly to minimize sample carryover and ensure a leak‑free flow path. Comparison of Agilent and third-party needle seat assemblies shows large differences in design. While the
other vendor still uses the older design, Agilent introduced a design in 2011 with more robust material, improved
performance, higher reliability, and larger pH range (0 to 13). The conical geometry at the center of the needle
seat also reduces sample dispersion. In addition, Agilent needles and needle seats are thoroughly tested to guarantee
full functionality for more than 30,000 injections.

Lifetime
Agilent rotors are rigorously tested and guaranteed for at least 30,000 injections. After 30,000 switch cycles, the Agilent rotor surface still seemed flat and consistent, and the contacting stator surface appeared clean. In
contrast, the third-party rotor already showed severe surface damage and a contaminated stator surface after 26,000
switch cycles. Therefore, shorter lifetime and potential carryover and leakage are expected when using
third-party seals. Photomicrographs showing the superior smoothness and integrity of the surface of an Agilent rotor seal.

Rotors
The rotor is a highly stressed part of the autosampler that is constantly switched back and forth, sliding over the stator. Its durability and life time is governed largely by the material and surface finish. Comparison of Agilent and third-party rotor seals revealed major differences in these aspects.

Surface smoothness
Microscopic inspection (Figure 9) shows the consistent flat surface of Agilent rotor seals, while scratches, flecks, and a jagged hole edge are clearly evident on the third-party rotor seal. The flecks indicate inconsistent material composition, and the scratches and jagged edge can definitely affect the sealing of the rotor to the stator, resulting in leakage or increased sample carryover.

Packaging
Scratches and flatness of rotor seal surfaces can also resultfrom poor packaging. Agilent rotor seals are packed in shapestable plastic boxes to avoid surface damage and deformation of the seal during storage and transportation, while third-party rotor seals are packed in normal plastic bags without special protection.

Reference
© Agilent Technologies, Inc., 2015
www.agilent.com/en-us/services/analytical-instrument-services/repairmaintenance/crosslab-preventive-

written by Ayodeji Ogunlowo

ayodeji@aasnig.com

Best Practice for Identifying Leaks in GC and GC/MS Systems (Part 2)

CONCLUDING PART

 Checking GC Connections

Checking all fittings for leaks immediately after installation, maintenance, and periodically while in use is an excellent practice. A handheld electronic detector capable of detecting a helium leak of 0.0005 mL/min in air is available commercially. Handheld leak detectors are particularly useful for finding leaks quickly either inside or outside the GC oven. It is good practice to always use a leak detector to check for leaks each time a column, fitting, or cylinder is changed. An excellent starting point for system troubleshooting is to first check for

potential leaks. Avoid using water soap solutions, as these can be drawn back into the GC flow path, severely impacting chromatographic results even to the point of causing permanent column damage.

Checking a GC/MS for Leaks

A vacuum or ion gauge, if ordered with your instrument, is useful for monitoring vacuum pressures under typical operating conditions in GC/MS. A vacuum gauge is very useful for isolating potential leaks to either the vacuum (MS) or pressurized (GC) side of the instrument. Vacuum readings in the 10–5 or 10–6 Torr range are typical for a system holding vacuum with a flow rate of 1.0 mL/min on a 30 m × 0.25 mm, 0.25 μm GC column. When the MSD is capped and pumped down, vacuum readings typically drop to the 10–6 or 10–7 Torr range in the absence of a leak. If the vacuum pump does not reach these levels relatively quickly, a leak somewhere in the MS is indicated. Make sure the purge vent is closed, the transfer line fitting is installed correctly, and that the large O-ring on the vacuum side plate is positioned correctly.

A software-based performance check of air and water is available in most GC/MS. This check looks at GC/MS ion traces of molecules typically found in air relative to ion 69 found in the calibrant. Ions 18 (water), 28 (N2), 32 (O2), 44 (CO2), and 69 (typical base peak from PFTBA used during auto-tune) are all monitored. Nitrogen (28) levels above 10% relative to the 69 peak indicate that the system has not had sufficient time to pump down or that there is an air leak. An air leak will typically show nitrogen:oxygen in a 4:1 ratio. Water (18) is also typically present, particularly after a system had been vented and exposed to ambient air. An equilibrated leak-free system should show nitrogen (28) well below 10% with oxygen 32 at approximately ¼ of the signal seen for nitrogen, and ideally water (18) lower than the N2 (28) peak.

Troubleshooting leaks in GC/MS is a process of elimination, looking at each site where a leak can occur. A fluorocarbon (for example, 1,1,1,2-tetrafluoroethane, ions 69 and 83) or argon (ion 40) spray can with a plastic tube to direct the flow is very useful in isolating a leak. A short spray at a suspect point and monitoring the appropriate ions in manual tune is a powerful tool for isolation. Key points to check are the transfer line connection in the oven, septum nut, column nut, and the large O-ring on the vacuum plate of the MS. Once a leak has been isolated the leak can be remedied by replacing a septum, resetting a column connection, or cleaning the O-ring on the vacuum plate, and reinstalling it back into the groove on the plate.

Innovations to Minimize Leaks

Figure 2 shows a total-ion chromatogram for an air and water check on a system that is operating normally. In this case, self-tightening column nuts were installed at the transfer line and inlet fittings. These column nuts provide a leak-free seal using a short polyimide/graphite ferrule at both column connections, without the need to retighten the fitting after more than 300 heat cycles. Use of these column nuts eliminates the need to retighten the inlet or mass spec transfer-line connections after oven heat cycling. Furthermore, because very low torque is needed to make a leak-free seal when using the self-tightening column nuts, these nuts are installed using only fingers, not wrenches, which eliminates the risk of over tightening and damage to the fittings (see Figure 3)

                                                                                                                                                                                               Figure 3. Agilent self-tightening column

     Figure 2. Example air and water check                                             nuts installed at the transfer line and inlet fittings.

Conclusions

By using tools, supplies and best practices that provide a leak-free GC or GC/MS, analysts can improve performance and productivity of their system. Agilent UltiMetal Plus Flexible Metal ferrules provide robust leak-free column connections, along with an inert surface for fittings in the sample flow path. The Agilent innovative self-tightening column nuts using standard short polyimide/graphite ferrules eliminate the need to retighten GC column fittings, including the mass spec transfer line, after repeated heat cycling. These new fittings also have the advantage of using only short polyimide/graphite ferrules for inlet, detector, and

mass-transfer-line connections. Following the best practices described in this technical overview and accessing the references below will help GC and GC/MS users identify potential air leaks, where to find them, and how to fix and prevent them quickly. One rule of thumb is to adjust fittings, septa, and O-ring seals to be JTE

for the best results

 CONCLUDED.

 References

Best Practice for Identifying Leaks in GC and GCMS:  Technical Note, Agilent Technologies

 

Written by Muyiwa Adebola

muyiwa@aasnig.com, www.aasnig.com

07084594001, 07084594004

Solve Carryover Problems in Gas Chromatography

Let’s first properly define carryover in the context that I’d like to discuss here. An injection is made and a chromatogram obtained. On injecting a “blank” as the next injection, one or more of the components of the previous injection appear in the “blank” chromatogram.

This definition needs further clarification:

  1. Blank injection can be a pure solvent (or solvent mixture), or, if contamination of the solvent(s) is suspected, then the “blank” may be an injection of air (for example, a 0 mL injection).
  2. While the “blank” may contain components of the previous injection, depending upon the solvent used, one may observe carryover from several injections previously. Here, I refer to the often puzzling issue where several injections are made without evidence of carryover then “out of the blue” a component will appear in the chromatogram that was present in a sample from several injections ago. I will explain this more fully subsequently.

In summary, carryover in this context is related to the instrument rather than solvent contamination. However, while the contamination may not arise from the sample solvent injected, the nature of the solvents used within the system are intrinsically linked to the majority of carryover problems in gas chromatography (GC). So, why is this and what can be done about it? To help with the visualization of the concepts discussed below, Figure 1 is a schematic of a typical split–splitless inlet with the various gas flows noted.

Injection Volume Related Carryover

In split–splitless injection, the injected liquid expands rapidly to form a gas plasma containing our analytes, hopefully also in the gas phase. The space within the inlet, into which this expansion occurs, is primarily dependent upon the internal volume of the inlet liner used, and the available volume may vary significantly depending upon the design of the liner used. Most manufacturers will publish the internal volume of their various liner styles or this information will be available through one of the many online vapour volume calculators: CHROMacademy Calculator (1), Agilent GC Calculator (2), and Restek Backflash Calculator (3).

The inlet pressure (determined by the total flow into the inlet), inlet temperature, and sample volume injected all influence the volume of gas created from the injected sample. If the volume of gas created by the sample exceeds the available volume within the liner, gas may overflow from the liner and will typically end up in the septum purge and carrier gas lines (yes, overflowing sample vapour may overcome the forward pressure of the inlet gas supply and flow back up the carrier gas inlet lines). As these lines are typically unheated, higher boiling and more polar sample components may condense and “coat” the lines. Any subsequent overloaded injection (typically known as “backflash”) may then flow through the unheated lines and re‑solubilize the condensed components. As the inlet pressure equalizes, or as the split line is opened in the case of splitless injection, then this re-dissolved component is drawn back into the inlet and may ultimately enter the column, therefore causing carryover.

One should note that this may not occur every time a backflash injection is made, and in some cases the polarity of the sample vapour relative to the condensed contaminant will determine how well the contaminant is re-dissolved, and therefore whether it is seen within the subsequent chromatogram. This can lead to the situation in which the contaminant may not become obvious until several injections later, if the solubility of the contaminant is not high in the intermediate injections. So, a polar contaminant may not appear in backflashed injections of, say, hexane, but may then become apparent when a backflashed injection of methanol is made several injections later. Not only is the problem of backflash rather insidious, it can also be very confusing because the appearance of the carryover may appear to be random and not apparent in the injections immediately following.

As splitless injections have inherently lower total gas pressure in the inlet (only the carrier flow is passing through the liner) and as the residence time of the sample solvent within the inlet is higher, splitless injection is considered to have a higher risk in terms of injection backflash and inlet contamination.

In order to overcome backflash issues there are three choices:

  1. Use pressure pulsed injection—in which the inlet pressure (total flow into the inlet) is increased during the injection phase and then reset to the desired pressure (and column flow) post injection. In splitless injection the pressure pulse time is usually matched to the splitless time.
  2. Reduce the amount injected
  3. Use a small split to increase the inlet pressure during injection

Obviously choices 2 and 3 need to be evaluated against any loss in sensitivity unless the sample concentration can be increased prior to injection, whereas choice 1 will
not lead to any reduction in analytical sensitivity.

The calculators mentioned above can all be used to assess the likelihood of backflash within the inlet and therefore will help to mitigate the issue.

Contamination of the Split Line

You may have noted that the split line on your gas chromatography instrument is also unheated. So whatever issues occur with the deposition of components into the carrier and septum purge lines may also occur within the split line. However, most inlet designs have a split line lower within the inlet than the split line (Figure 1), and the sample gas typically follows a more tortuous path through the liner prior to passing out of the split line. This being said, it is perfectly possible for less volatile sample components to condense within the split line and the charcoal trap, which is also included in the split line on most instrument designs. If the concentration of this contamination is high or builds up over a period of time, it is possible for carryover to occur in a very similar fashion to the backflash injection. This can be confirmed, and indeed mitigated, by “steam cleaning the split line” with several large volume injections (typically 5 mL) of water at very high split flows. If the contamination is nonpolar, then ethyl acetate can also be injected in a similar fashion until the carryover is eliminated.

Contamination of Inlet Components

Inlet components such as the liner may become “active” over time, as a result of exposure of silanol groups on the quartz glass from which the liner is made or from any quartz wool packing within the liner or from active metal sites on the inner metal surfaces of the inlet body. This is typically associated with peak tailing phenomena for polar analytes because of unwanted secondary interactions between the analyte and the active inlet site. However, if this interaction is strong, sample components may be irreversibly adsorbed until the following injection. However, again, if the solvent used for the next injection does not readily dissolve the contaminant, the carryover may not occur until an injection of a solvent of the same polarity is made.

To avoid these issues, ensure that deactivated liners are used, avoid the use of glass wool and packing materials within the liner if possible (check the impact on analytical sensitivity and reproducibility and discrimination effects before moving to a liner with no packing), and ensure that the inlet body is regularly cleaned. Liners should be regularly replaced and should be changed as a matter of priority when carryover problems are being investigated.

Carryover from the underside of the septum can also occur if the septum purge flow is not sufficient. This gas flow is designed to flush away septum outgassing products and to avoid sample deposition on the underside of the septum. Many instrument designs have automated control of septum purge flow and so this variable is rarely considered or the flow manually measured. Septum purge flow should be manually measured as part of preventative maintenance routines.

Contamination of Autosampler Components

Finally, the syringe and wash solvents should be considered in any investigation into carryover. Contamination may be carried on the inside and outside surfaces of the syringe needle and this can be somewhat mitigated by using rapid plunger depression and very short residence times for the syringe needle within the inlet. Most manufacturers ensure that this is built into the autosampler routine, however if your instrument provides the option for “slow” or “fast” injection, be sure to choose the fast injection option, especially with splitless injection.

Further, the syringe wash solvents should be matched to the polarity of the potentially contaminating analytes. While most users match the wash solvent with the sample solvent, one needs to carefully consider the solubility–polarity of the components involved in carryover when selecting the wash solvent. Further ensure that waste solvent bottles and the bottle tops are kept clean and regularly emptied. When the autosampler offers more than one wash solvent, one should experiment with the wash solvent routine to minimize carryover—use of one solvent post injection and one solvent pre injection, or the use of both solvents in turn both before and after injection, and so on.

The number of sample washes and syringe primes prior to injection can also be optimized to reduce carryover to the minimum levels. I have known methods that will be carryover free only after five sample washes and five sample primes prior to injection.

Hopefully you will now have a more thorough understanding of the potential sources for carryover that are instrument related and that some or all of this advice will help you to overcome issues with quantitative reproducibility or contaminants in qualitative analysis.

References

  1. https://www.chromacademy.com/GC-Sample-Introduction-Best-Practice.html
  2. https://www.agilent.com/en/support/gas-chromatography/gccalculators
  3. http://m.restek.com/images/calcs/calc_backflash.htm

Akinbuli Opeyemi

www.aasnig.com, opeyemi@aasnig.com

08068129603.

Simple Steps for Clearing a Blocked Injector in Your ICP-OES Torch

Remove blockages to ICP-OES productivity

Deposition of the sample matrix, salts or even carbon build-up can lead to injector blockage in the torch. How quickly blockages occur varies, depending on sample type, sample workload, torch type, and even the method parameters. A blocked injector can restrict the flow of sample aerosol into the plasma, decreasing sensitivity and
degrading accuracy and precision.
Prevention is the best cure to reduce injector blockage and extend the operating time. Make sure that you are using the recommended torch type and check that you have the recommended instrument parameters for your application. Filter all samples to ensure you remove large particulates. Regular rinsing between samples and at the end of the run can also help to keep the injector clear. However, improper cleaning techniques can permanently damage the
torch. Follow the steps outlined in this technical overview to safely clean your torch, and to remove blockages if or when they occur.

Simple Steps for Clearing a Blocked Injector in Your ICP-OES Torch

Routine cleaning


5100/5110 ICP-OES
– Prepare a 50% aqua regia solution (1 part deionized water to 1 part aqua regia [three parts hydrochloric acid and one part nitric acid]) in a wide diameter tall form beaker.
– Place the beaker under the torch cleaning stand (P/N G8010-68021). This suspends the torch (or injector/base assembly) in the cleaning solution, reducing the risk of spills and damage to the quartz outer tube.
– Invert the torch and position this on the torch cleaning stand so that the quartz outer tube and injector is immersed in the aqua regia solution.
– Pipette some of the aqua regia through the ball joint of the injector to remove buildup from the lower part of the injector.
– Soak the torch for at least 1 hour.
– If deposits remain, repeat the cleaning process using a higher concentration of aqua regia.
– Thoroughly flush the inside and outside of the torch with deionized water (18 MΩ cm) using a wash bottle. Invert the torch and flush deionized water through the quartz tubes so that the water flows out of the gas entry ports and ball joint connector for at least 30 s.
– Invert the torch and dry by blowing clean compressed air or nitrogen through the gas ports on the base and through the opening of the ball joint to remove moisture.

Caution: Do not place the torch in a drying oven. It is not as effective at removing moisture as using compressed air or nitrogen, and may damage the torch.

One-piece quartz torch for 700, Vista, and Liberty Series ICP-OES

– Soak the torch overnight in concentrated aqua regia (three parts hydrochloric acid and one part nitric acid).
– If necessary, use a pipe cleaner dipped in aqua regia to gently remove persistent deposits from the injector tube.
– Rinse with deionized water and allow to dry.

Removing salt deposits
– Rinse the torch with water.
– Soak overnight in a 25% detergent solution.
– Rinse the torch with deionized water and allow to dry.
Important: The torch must be completely dry before re-installing. Replace the torch if chipped, cracked,
or distorted.

Re-installing the torch for 700, Vista, and Liberty Series ICP-OES

– Position the torch in the center of the RF coil, resting on the torch stand.
– Close the torch clamp and turn the locking knob.
– Gently attach the transfer tube to the base of the torch.
– Align the torch so the intermediate tube is about 2 to 3 mm away from the RF induction coil.
– Connect the auxiliary and plasma gas hoses to the appropriate inlets on the torch.
– Complete the torch alignment procedure to ensure the optics are viewing the highest emission signal from the plasma.

Caution:
– Never place a torch in an ultrasonic bath, or use a wire to clean the injector.
– Do not use hydrofluoric acid with glass or quartz sample introduction components.
– Always use care when handling or installing a torch. Excessive force can break the torch.
– Do not touch quartz torches with bare hands. This can
reduce torch life.

– A one-piece quartz torch is simple to install and use and delivers great performance for most applications.
– For organic solvents, use a torch with a smaller ID injector. For volatile organic solvents, use a torch with a narrow-bore (0.8 mm ID) injector.
– For greater flexibility and reduced running costs, choose a semi-demountable torch. The injector and/or outer tube
is removable and replaceable separately.
– For fusions and HF digests, use a torch with an alumina injector.
– A fully demountable torch allows you to replace components individually, which can help lower your operating costs.

Learn how to achieve better sensitivity and precision, and improve tolerance to samples with high levels of total dissolved solids (TDS) by switching to the Agilent OneNeb Series 2
nebulizer.

written by Ayodeji Ogunlowo

ayodeji@aasnig.com

A Guide to Transporting Materials

It’s one thing to ensure your lab is operating safely, but the regulations and processes surrounding the transportation of hazardous materials are especially stringent. Whichever method these materials are being transported by, their packing and transportation must adhere to international regulations.

Safety is of the utmost importance in this process. In this guide, we’ll talk you through the differing material classifications, the regulations surrounding materials and modes of transport, as well as any necessary training required for transporting chemicals.

Types of materials and classifications

When processing, packaging or transporting dangerous goods, you will need to be able to classify them correctly so everyone involved in the process of transportation is aware of the potential hazards.

Based on their main hazards, goods are assigned to different numbered classes, as decreed by the UN and are as follows:

UN Class

Dangerous Goods

Classification

1

Explosives

Explosive

2

Gases

Flammable gas; non-flammable, non-toxic gas

3

Flammable liquids

Flammable liquid

4

Flammable solids

Flammable solid; spontaneously combustible substance; substance which emits flammable gas in contact with water.

5

Oxidisers and organic peroxides

Oxidising substance; organic peroxide

6

Toxic and infectious substances

Toxic substance; infectious substance

7

Radioactive materials

Radioactive material

8

Corrosive substances

Corrosive substance

9

Miscellaneous dangerous substances

Miscellaneous dangerous substances

Regulations covering different materials and types of transport

Whether you’re transporting materials in the UK or throughout the European Union, your legal obligations are broadly similar and must adhere to the respective regulations governing transport by road, rail, inland waterway, sea and air.

By road: If you’re transporting by road, you must adhere to the International Carriage of Dangerous Goods by Road Regulations, while domestic transportation must stick to the Carriage of Dangerous Goods and Use of Transportable Pressure Equipment Regulations.

By rail: Governed by Appendix C of the Convention Covering International Carriage by Rail, train transport must adhere to the regulations covered within.

By sea: When transporting by sea, you must adhere to guidance provided by the International Maritime Dangerous Goods code, which is used by operators transporting dangerous goods that need to travel across seas.

By air: The International Civil Aviation Organisation’s Technical Instructions are an internationally-agreed set of provisions that dictate the requirements for transporting dangerous goods by air.

Required training for transporting chemicals

If your business is frequently involved with the handling, processing or transporting of dangerous goods then you must appoint a Dangerous Goods Safety Adviser as per the Health and Safety at Work Act 1974. A DGSA is not required if you transport smaller quantities of dangerous goods than specified in the legislation, or your business is only occasionally involved in the transportation of dangerous materials.

The DGSA is required to monitor your business’ compliance with the rules that govern transportation, provide advice on said transportation and prepare annual reports on the business’ activities in material transportation. They are also responsible for monitoring safety measures, investigating accidents and advising on the potential security aspects of transport.

It is mandatory for DGSAs to obtain a vocational training certificate upon completing the correct training, which involves passing a written examination. The training courses for DGSAs are run by independent providers and the trade associations for each mode of transport, and vary in length from two to five days, depending on the mode of transport covered.

Minimising risks during transportation

Whichever mode of transportation is being used, there are a number of risks that could occur in transit, including damage, theft, chemical burns, fire, explosions, as well as many more. Consider the following before you make your journey:

  • Make sure large or heavy loads are protected properly and in the appropriate manner
  • Very important: ensure weight is evenly distributed.
  • You may need goods-in-transit or marine insurance to protect goods being transported, which may be paid for by the buyer or seller of goods.
  • Put suitable warning signs on vehicles to indicate wide, long or hazardous loads.
  • Take the appropriate security measures. For example, high-value goods should be tracked using a vehicle-tracking system.
Reference
www.mynewlab.com

Posted by Adi Oluwakemi

kemi@aasnig.com, www.aasnig.com

08060874724, 07084594001

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