Category Archives: How-To & DIY

UV-Vis Spectrometry – Troubleshooting & DIY

UV-Vis Spectrometry

UV-Vis spectrometry is a simple, but sensitive technique that’s most often used to quantify, and less commonly used to identify. The concept is straightforward: light goes in, and less light comes out. Spectrophotometry is so attractive because of the technique’s sensitivity: its detection is linear across three orders of magnitude. Nearly any sample concentration can be determined by choosing an appropriate dilution. By applying the Beer-Lambert law, analysts can quantify the amount of a chemical present in a sample with very high precision.

Beer Lambert Law © VerdantThe Beer-Lambert Law

By measuring the amount of light that didn’t make it through (absorbance, A). Every chemical has a unique physical property called an extinction coefficient, which is a measure of how much light it absorbs (ε = color intensity in solution). This value is specific to a given wavelength, so this is sometimes written with a subscript, such as ε690. When you combine that information with the amount of sample the light passed through (l – the path length), you can determine an exact concentration (c).

Important! Wavelengths are usually chosen by the absorbance maximum (highest peak), but in advanced cases, different wavelengths can be used to bring the sample’s absorption into the linear range without dilution.

What’s the difference between spectrometry and spectrophotometry?

Not much – the difference is what you do with the information. Spectrometry is evaluating a profile of electromagnetic radiation for a compound across an entire spectrum. That can be an entire range, such as all UV & visible light, or it can be a limited subset like the near infrared. A UV-Vis spectrum can be used to identify compounds in a sample, or tell inorganic chemists about the nature of the bonds in a complex. Spectrometry applies to any part of the entire electromagnetic spectrum.

Spectrophotometry deals with measuring the amount of light being transmitted or absorbed at any point along that spectrum. That enables an analyst to to determine the concentration within a sample, and provides researchers with important information about a sample’s physical properties.

Important! Light is measured in moles of photons. Absorbance is a unitless measurement because the moles cancel during calculation – another name for ϵ is the molar absorption coefficient.

Proper Maintenance


Troubleshooting UV-Vis Spectrometry

I don’t see any peaks! (Flat Spectrum)

First, ensure that your lamps are turned on, warmed up, emitting light, and that the light path to the detector is not blocked.

If your UV-Vis uses a monochromator instead of a polychromator & photodiode array, it’s possible that the motor has failed or become stuck. If this is the case you will have a flat graph as it reads every data point to be the same, but it will be offset from the baseline to some extent when running a sample compared to a blank. To confirm, compare a blank to a sample covered in electrical tape, which be read as a sample with a saturated absorbance signal. Motors fail often are not too hard to replace, but this will be a precision servo. Replacing it in the proper alignment can be challenging.

If you can visibly confirm that your lamps are turned on and working, then you may have a problem with the detector. If you have a working monochromator, it’s reasonable to expect that a single photodiode may have failed. But in the case of a polychromator and photodiode array, it’s very unlikely that an entire array failed at once. Trace the wiring inside the instrument and to the computer, checking for loose cables, unseated connectors, or broken/sharply bent wires.

I don’t see any peaks! (Noisy Spectrum)


My spectrum is flat on the left or right side! (Spectrum Trails Off Flatly)

If your spectrum begins to flatten toward the left or right side, it means you have a dying lamp. If it’s toward the left side, in the UV range (~200 nm), then it’s your deuterium lamp. If it’s to the right side, in the red light range (~700 nm), then it’s your tungsten lamp. Both lamps emit overlapping wavelengths of light in the center of the spectrum, so that’s why you’ll only see dead zones in the far ends are specific to each lamp.

My spectrum is flat on the left side! (Noisy Left Side)

You’re most likely using the wrong type of cuvette. Plastic cuvettes won’t work because most polymers are linked by a carbon-carbon single bond, which absorbs strongly in the UV range. Fused quartz cuvettes are used because it’s one of the few substances which allows ultraviolet light to pass absorbs ultraviolet light.



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Infrared Spectrometry (IR/ATR) – Troubleshooting & DIY

Troubleshooting Infrared Spectrometry (IR/ATR)

Developments in ATR have caused it to become the most common form of IR spectroscopy over the past two decades. Its unparalleled convenience has led to widespread use in preliminary and routine analyses. But the deceptive simplicity of ATR can lead to some common, but unexpected issues.

Although ATR is attractive because it requires little to no sample prep, be aware that the prep work has just been shifted to the instrument itself – and is still as important as ever. Since your sample’s spectrum will only be as good as your contact with the crystal – your first step will be proper cleaning.

Prep Work & Preventative Maintenance

The ATR crystal should be cleaned thoroughly at the beginning of every work day, and after any analyses which might leave a residue. These can include foods, oily products, plasticizers, and objects exposed to the outdoors. First, wipe the lens and surrounding area using lint-free optic lens cleaning paper, such as Kimwipes. Next, remove any leftover residue with a fresh wipe soaked in an organic solvent such as methanol or MEK (the higher the volatility, the better).

Important!  Water should never be used! Although some crystals can tolerate exposure, there’s always a better solvent choice. It has high IR absorption, evaporates slowly, and avoiding water altogether will remove the potential for expensive mistakes. Acetone is suitable, but should be avoided – stray droplets will mar the instrument’s plastic surfaces.

After cleaning the surface with a solvent, it is very important to allow for full evaporation before taking a background. If the solvent doesn’t fully evaporate first, the spectrum will be skewed by the solvent peaks. If the ATR window is under an argon or helium gas purge, the close the gas valve and open the chamber to room air. Nitrogen purges can remain in place and will be fully evaporated after ten minutes. Evaporation in standard room air takes approximately 20 minutes. Permanently enclosed chambers require even longer.

Important!  Solvent evaporation depends on its attraction to molecules in the air. Inert purge gases like helium and argon simply don’t have the intermolecular attraction to carry the solvent away at a reasonable pace.

Dry and powdered samples should be cleaned after every use by wiping the area thoroughly with lens cleaning paper. Follow this by using a non-abrasive powder brush to dislodge any stubborn particles which may still be caked onto the sides of the crystal. Once the instrument is clean, double check your method to ensure that it’s set up properly for your source and ATR window. Ensure that you have a reasonable scan resolution and count, and take a new background. Then perform a scan of a standard such as PS, PTFE, or LDPE film. Compare this to a reference spectrum.

Important! If your peaks are shifted from the correct wavenumber, your instrument could need realigned or recalibrated. If your peaks are growing weaker than your reference spectrum, your source could be aging; the ATR window might need polished/replaced; or the clamp could be giving poor contact with the window.

I’ve got this weird band around 3500 cm^-1 that keeps creeping in every few days. I can’t figure it out!

Important! That’s part of the water spectrum – you’re seeing the change in humidity. Controlling the environmental conditions in your lab is an topic unto itself, which we’ve explained in detail here.

If your IR is cooled or purged by gas from a liquid nitrogen tank, condensation is also a possibility.

My spectra used to be smooth but now they’re jagged and I don’t know what happened!

Important! Most likely, the scan resolution was changed somehow. This can happen when a different method gets loaded. Most labs run their instruments at low resolutions for smooth graphs with easily identifiable peaks. Higher resolutions will suffer a signal-to-noise ratio drop. To maintain the same quality in resolution half as wide, you need to double your scan number (and 1/4 width requires 4x the number of scans; and so on). This quickly leads to long sample collection times for instruments which aren’t designed for high-res data collection.

What’s a sine wave doing in my data?

Important! Your sample is probably reflective. If the problem persists through different samples, then a mirror within the instrument may have fallen out of position.

I’m getting results that don’t make any sense!

Important! Spectrum database search software often gives questionable results, especially when analyzing composite / proprietary samples. Create a table of molecular features from the peaks and bands; and supplement your analysis with data from other instruments when needed.

I’ve got a massive backlog of samples to identify or compare. Have you got any tricks that can help me? 

Important!  Try rapid qualitative analysis to quickly identify samples of interest:

First, configure your IR method with a resolution and scan count that produces an acceptable spectrum in less than five seconds. Next, clean the window with solvent, wait 60 seconds, and immediately perform a background. Then run each sample exactly 60 seconds after cleaning the previous residue from the window. The residual solvent peaks will be suppressed by the background. Repeat samples of interest using proper techniques, as these spectra will not be suitable for publication, consumer goods, or physical evidence.


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Thermogravimetric Analysis (TGA/DTA) – Troubleshooting & DIY

Troubleshooting Thermogravimetric Analysis / Differential Thermal Analysis (TGA/DTA)

Important! This page is under active construction – please check back in a few days. Important!



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Differential Scanning Calorimetry (DSC) – Troubleshooting & DIY

Troubleshooting Differential Scanning Calorimetry (DSC)

Important! This page is under active construction – please check back in a few days. Important!



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Liquid Chromatography & HPLC – Troubleshooting & DIY

Liquid Chromatography & HPLC

Liquid chromatography is one of the most common processes in science. You may you know it as HPLC, and use it for analysis. Or perhaps you know it by a trade name, such as CombiFlash, and use it for separation of products. Whatever the situation, it’s rare to find a lab that doesn’t employ some form of liquid chromatography. In a sentence, LC involves passing a liquid through a tube of granular material that will interact with the samples dissolved within the liquid. The tube is called a column, and its solid contents are the stationary phase. The liquid that passes through the column is the mobile phase, called an eluent or solvent. The interactions between the stationary and the mobile phase happen at the molecular level – but (ideally) there is no reaction or filtration taking place.

LC Column

LC columns are classified as either normal or reversed phase. A normal stationary phase is neutralized silica gel (essentially high purity sand with its pH adjusted to around 7). Other varieties include silicate polymer gels bound to other polar groups, such as nitros or diols. Reversed phase consists of a hydrocarbon gel categorized by its chain length or monomeric sidegroup (C18, C8, phenyl, and so on). Column type is your first consideration in LC, and it will mainly be influenced by your sample’s molecular properties.

  • Reversed Phase (RP – Nonpolar): Hydrocarbon or aromatic polymer.
    • Interacts with non-polar compounds, slowing them down (longer retention time).
    • The most polar compound elutes first; most non-polar compound elutes last.
  • Normal Phase (NP – Polar): Neutralized silica or silicate polymer with polar side-group.
    • Interacts with polar compounds, slowing them down (longer retention time).
    • Most non-polar compound elutes first; most polar compound elutes last.
  • Special Types
    • Ion Exchange: Polymer gel with ionized side-groups. Reacts with specific ions in sample to quantify or purify.
    • Size Exclusion: Dextran polymer gel network. Hinders sample passage to separate compounds by molecular weight.

Important!  Polymers are less physically stable than silica-bonded columns. Always check your method against the maximum pressure specifications of the column.

Eluent (Solvent)

Your mobile phase selection will depend on your sample composition and available columns. Solvent quality is extremely important for LC. Impurities accumulate on the column and impede flow. Dissolved gases can lead to measurement error if not removed.

Protic solvents normally involve the use of a buffer solution to maintain the column at optimal pH, but these pose a risk to the instrument itself. Since salts and ionic solutions are corrosive, these must be purged from the system when not in use.

Important!  Never transition directly from a buffered solvent to a storage solution such as acetonitrile! Always flush buffers with water or another protic solvent first! Otherwise the salts will precipitate into the tubing or column fittings blocking flow and causing localized corrosion.

Verdant is an industry leader in solvent recovery technology. Recycling used solvent is a key focus of industrial sustainability efforts – and it’s also a great way to reduce operating costs. If you’re interested in learning about your options, ask us how we can help.

Flow Rate

Your instrument’s flow rate affects the pressure within the system. Your flow rates will typically fall within a range varying on the order of milliliters per minute (mL/min). Your flow rate selection will be influenced by your column dimensions, pore size, stationary phase composition, and solvent viscosity.

Data Sampling Rate

If your LC has a variable data sampling rate, faster is better – up to a point. If your sampling rate is too low, narrow peaks will tend to clip – meaning you’ll miss the tip of the peak and get an inaccurate reading. Set your data sampling rate to 40 Hz as a starting point, and then raise it as high as you can without increasing baseline noise.

Derivatization / Internal Standardization

Derivatization is an excellent option when you have trouble with solubility, detection ranges, or peak shape. By subjecting the sample to a chemical reaction, you can change the chemical structure into a derivative that’s better suited for analysis. It can also be used to work with a more limited set of  supplies by increasing sample compatibility with your available columns and solvents. There are more reactions than we could ever list here – entire books have been filled with the possibilities. But don’t worry, because we have those books! Contact us for assistance if you need help with this technique.

Internal standardization is a technique that’s better suited to UV detectors than LC-MS. This can help you to determine something elusive that can’t normally be detected with UV – such as a sample and solvent pair that absorb in the same range. By spiking your sample with a compound which absorbs outside of that range, you can quantify concentrations much more accurately.

Troubleshooting Liquid Chromatography & HPLC

I have a noisy baseline! (General Issues)

A noisy baseline is a common symptom with several different potential sources, which can make troubleshooting very time consuming. You might have a leak. The column could be contaminated or degrading. You could have a low signal to noise ratio brought on by solvent impurities, or a data acquisition rate that’s too high for your method. If the noise is periodic, or rhythmic like a wave, then it could be a failing pump or pulse dampener. If the noise is spiky and appears irregularly, it there could be bubbles within the system, or the detector lamp could be flickering/failing. If your instrument isn’t isolated on a power filter with ferrite chokes on the wires, it could also be caused by electrical noise from other devices within the lab.

My baseline won’t stay still! (Baseline Drift)

If your baseline repeatedly drifts in the same direction over the course of every gradient run, then your column is contaminated (if you normally run isocratic, you can run a gradient to see whether this is the case). Ensure that you’re taking adequate column protection measures, then clean or replace your column.

If you’ve ruled out column contamination, or if your baseline is drifting randomly or steadily in a particular direction over the course of multiple runs, it may be time to replace your detector lamp.

My peaks keep showing up earlier/later every time I run a sample! (Retention Creep)

As your column ages, it will be less effective at interacting with the sample, and the retention time will slowly get shorter. If this effect becomes significant enough to be noticeable between runs, check that you’re adequately maintaining the column’s pH between runs. If so, then it’s just time to change the column.

If your retention time is getting longer with each run, remember that LC leaks cause late peaks. If you can see the problem worsening, then you likely have a part wearing out and physically falling apart, which usually be accompanied by other erratic findings. You’ll most likely find the offending part, but LC is a sensitive technique, so don’t be surprised if the flaws aren’t visible to the naked eye. When in doubt, replace any parts which have been in service long enough to be at risk.

If your retention time is changing erratically, there is probably a bubble entering the system at injection time, or solvent cavitation occurring at the pumps. Ensure that your sample solvent and mobile phase are being adequately degassed, and check the inlet.

My peaks have peaks! (Split Peaks)

This means your sample is getting hung up on something. First, confirm that your sample and its solvent are compatible with your entire mobile phase (both solvents if you’re running a gradient). If compatibility checks out, then there is something in the way. This can either be a solid contaminant, or a bubble trapped in a component. Careful inspection will be necessary to determine which.

My peaks aren’t sharp enough to quantify! (Peak Tailing / Fronting)

This usually indicates a problem with the column caused by improper use. First, rule out whether your sample volume is overloading your column. Then check the column for voids, degradation, or channeling (all ways of saying: gaps within the column packing). For troubleshooting this issue, the simplest first step will be to replace the column and guard. Before running another sample, check for flawed processes. Confirm that the method is not exceeding the temperature limit of the column packing. Ensure that the column’s pH is being maintained correctly. Verify that the column is being properly stored between uses. Consider implementing a log recording the dates the guard columns are replaced, and increase the frequency if necessary.

If the problem is unique to particular samples, or certain portions of the chromatogram, you’re most likely seeing multiple components elute together, and should adjust the method to secure better separation.

My peaks are too wide! (Column Capacity)

Issues with peak width are mainly symptoms of sample volume. If your peaks are too broad, your column is most likely too small for your sample size, and is being overloaded. This is especially true for complex samples – your column interacts with everything being eluted, not just your sample. Ensure that you’re factoring the entire sample into column size determinations. If you’re certain that isn’t the case, first make sure your fittings are secure and that your target pressure is being reached. Then try slowing your gradient transition rate down to allow for a slower elution time.

My peaks are running together! (Poor Resolution)

This is usually a sign that it’s time to replace your guard column or filter, particularly if you’re noticing that pressures are beginning to rise. This is because poorly soluble contaminants tend to build up at the front of the column and get stuck.

My analyte isn’t showing up! (Missing Peak)

One possibility is that your analyte isn’t compatible with your solvent or column choice. This can include insolubility, UV absorbance masking, or samples that interact heavily with the column.

Another possibility that the sample is too dilute. A UV detector is linear across 4 orders of magnitude (104, or 10,000x concentration difference), so try increasing the concentration if the dilution factor is within your control.

Nothing is showing up at all! (No Peaks)

This could be an issue with your detectors or a wiring fault preventing communication. If you see noise in the baseline, the issue is probably not a loose connection or faulty wiring. Verify that your detector wavelength is set to a value that’s compatible with your solvent & analyte, and confirm that the connected solvent is what you intended to use. Check the signal gain and attenuation to ensure that your signal isn’t being suppressed into the baseline. Next check and re-zero or replace your lamp. If the problem still persists, then your detectors may be at fault.

I see peaks I don’t expect / the same peaks every run! (Ghost Peaks)

First, identify your ghost peaks by running your gradient method without any analyte. You can minimize ghost peaks by using HPLC grade solvents, and degassing, filtering, or using purification/water removal technologies.

Next, identify any injection issues by injecting a blank sample. Problems linked to injection indicate a mechanical part failure. You may have a poor seal that’s leaking, or you may have pump seals or injection rotors breaking down.

If your ghost peak is inconsistent, you might have a late bloomer in your sample. You can check for this by running your last sample and doubling the method time. The best thing you can do is to flush your injector between samples, and incorporate a “Solvent B” purge into the end of your method.

There’s something wrong with this LC Injection Syringe!

Even carefully maintained and properly used syringes wear out quickly with normal use, so it’s best to consider your syringes as consumables. Injection syringes are extremely fragile, and prone to failure at the slightest misuse. LC can involve corrosive liquids which can jam up a syringe pump within a few exposures.

Proper cleaning includes flushing the syringe with an appropriate solvent after every sample. Follow this with water and several acetone rinses, and then place the syringe back into its case.

We recommend that you always have spares on-hand. Replacement parts can be helpful, but undetectable flaws can make it difficult to change them without accidental damage to the new parts. It doesn’t take many bent replacements before it becomes cheaper and less frustrating to simply have an extra.

I heard you can regenerate a used column. How does that work?

As you run samples and solvents through your column, it will become less effective. While it is technically possible to regenerate a column, it isn’t always helpful to try. So before we delve into the process, let’s take a look some situations where you probably shouldn’t bother:

:(  If you’ve run samples with no filter or guard column
:(  If the column has already been regenerated several times
:(  If the column’s connections or fittings appear visibly worn
:(  If the column’s packing has begun to ooze out or channel (gaps within the material)

If any of those conditions are true, then regenerating the column won’t be your best option. Column regeneration involves the use of several different solvents which your lab may not carry – this can be a large initial expense, take up valuable lab space, and add to the regulatory overhead. The potential savings may not be worth the time investment for busier labs.

Important!  If your lab typically runs samples of unknown identity, your components probably have very short lifespans. But this doesn’t have to be the case! You can safeguard your column by pre-filtering your analyte. A standard vacuum filtration through a Nylon 66 membrane filter will remove anything that would get trapped on the column or in your instrument fittings. There’s no need to worry about sample loss, because anything trapped by this filter wouldn’t have been suitable for LC analysis in the first place. You can also pre-filter your solvents if you have concerns over their purity.

It’s also important to have a realistic expectation of what column regeneration can offer. When done properly, it will extend your column’s useful life and remove ghost peaks, but it won’t bring your column back to 100%. A regenerated column is like a tuned-up used car: it may run perfectly fine, but there will always be an increased of developing problems.

The table below provides the steps for the most commonly used columns. Different processes are required for special columns such as zirconia, or columns used for proteins and other biological samples. You also can design a customized process suited for the solvents and columns used within your lab.

Regeneration Processes for Silicate Columns

Column VolumesNormal PhaseReverse PhaseIon Exchange
20Hexane60 °C DI Water60 °C DI Water
20Methylene ChlorideAcetonitrileAcetonitrile
2060 °C DI WaterHeptaneMethylene Chloride
20Mobile Phase/Solvent AMobile Phase/Solvent AMobile Phase

Important! Hexane can be substituted for any HPLC grade hydrocarbon solvent. Isopropanol can be substituted with any anhydrous, HPLC grade alcohol, but it is the best transition solvent, but it is very viscous and should be run at a moderate flow rate.

Important! If your contaminants are metallic, these can be removed by including ETDA in the DI water pass.

Important! We don’t recommend “all in one” column regeneration mixtures, because no mixture of solvents can properly treat the column packing in one step.

Column volumes are the number of times the void volume for your column must completely transit the column to achieve the intended result. In plain english: For every milliliter per minute of flow rate (r), a full equilibration will take twenty times your column’s void volume (v). In an equation: 20 × (v mL) × (r mL/min) = # of mins.

The void volumes for most standard column sizes are given in the table below.

HPLC Column Void Volume (0.70 Average Pore Volume)

Column Size (mm x mm)Void Volume (mL)
300 x 50412.33
300 x 2172.74
300 x 1016.50
300 x 7.810.03
300 x 4.63.49
300 x 2.10.73
300 x 1.00.17
250 x 50343.61
250 x 2160.61
250 x 1013.75
250 x 7.88.36
250 x 4.62.91
250 x 2.10.61
250 x 1.00.14
150 x 50206.17
150 x 2136.37
150 x 108.25
150 x 7.85.02
150 x 4.61.75
150 x 2.10.36
150 x 1.00.08
125 x 50171.81
125 x 2130.31
125 x 106.87
125 x 7.84.18
125 x 4.61.45
125 x 2.10.30
125 x 1.00.07
100 x 50137.44
100 x 2124.25
100 x 105.50
100 x 7.83.35
100 x 4.61.16
100 x 2.10.24
100 x 1.00.06
50 x 5068.72
50 x 2112.12
50 x 102.75
50 x 7.81.67
50 x 4.60.58
50 x 2.10.12
50 x 1.00.03
30 x 5041.23
30 x 217.27
30 x 101.65
30 x 7.81.00
30 x 4.60.35
30 x 2.10.07
30 x 1.00.02
15 x 5020.62
15 x 213.64
15 x 100.82
15 x 7.80.50
15 x 4.60.17
15 x 2.10.04
15 x 1.00.01

If yours isn’t listed, you can determine its void volume for your column by calculating its volume and multiplying it by the average pore volume (usually 0.70, often 0.50). Convert all column dimensions to centimeters to get an answer directly in mL (1 cubic centimeter ≈ 1 milliliter), and divide the column’s inner diameter by 2 to get the radius!  v0 mL = 0.7 × π × l cm × (r cm)².

Important! Several manufacturers have copied mistakes from each others’ materials, leading to widespread confusion about void volumes. The volume of a cylinder is calculated from its radius squared – don’t square the diameter!

As always, we encourage you to contact us if you need help.


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Gas Chromatography & GC-MS – Troubleshooting & DIY

Optimizing Gas Chromatography & GC-MS

GC-MS has become the gold standard for routine chemical analysis and definitive identification. But it has reputation as a time consuming technique, and lab productivity is commonly bottle-necked by the GC. Fortunately, you don’t need to purchase a second GC to increase your throughput, because there are several ways to speed things up.

Optimizing your GC method can save you hours each day, so it’s important for your lab to get the most out of your equipment. A stock GC configuration with a default method can take 40 to 60 minutes. But a fifteen minute run-time is a realistic goal for most labs.

Labs that analyze the same few compounds every day – like production labs – will be nearer to ten minutes. Labs which only perform analyses within the same category – such as research labs – will be closer to fifteen minutes. Labs which need to cover a broad range of possibilities – typically forensics labs – will have the longest times, but even these should complete in under twenty minutes.

Now that you know what’s possible, let’s talk about how to get there!

Carrier Gas

The three main choices of carrier gas are helium, nitrogen, and hydrogen. Each choice has certain advantages and disadvantages, but the main points to consider are cost, safety, effectiveness, and speed.

Nitrogen is the most commonly used gas. As the most abundant gas in the atmosphere, Nitrogen’s main advantage is that it’s the cheapest carrier gas. Its biggest disadvantage is speed, so it tends to work best in labs where the GC sees only occasional use but doesn’t run constantly. That slow pace is the reason nitrogen has the best column efficiency, so it can be a good choice for tough separations – but you can achieve the same result by optimizing other components. So when the lab-work is being is held up at the GC, the first thing you consider should be switching to a different carrier gas.

Of the three gases, Hydrogen offers the greatest speed advantage. It comes with a reasonable cost, and its physical properties make it well-suited for use across a wide range of speeds, which gives it greater flexibility. Hydrogen can function well equally well anywhere from 20 to 80 cm/s. By comparison, nitrogen’s column efficiency drops off sharply at any velocity except 10 cm/s. This versatility allows for the fastest separation of many different compounds with methods optimized for the target analyte. Hydrogen easily stands out as the best overall choice for a carrier gas, but its primary disadvantages are safety concerns.

Important! Hydrogen is a widely-used carrier gas with virtually no risk to trained professionals when properly implemented. But proper implementation does require an initial investment in specialized equipment. The greatest risk is posed by using a tank of compressed hydrogen. By opting for a hydrogen generator, the gas is never pressurized and the risk of self-ignition is removed. The remainder of the risk can be mitigated by using a nitrogen purge or hydrogen detector to guard against leaks.

Important!Important! We advise against hydrogen as a carrier gas in academic labs where introductory training occurs with limited supervision.

Helium represents something of a compromise between nitrogen and hydrogen. It has a significant speed advantage over nitrogen; it’s almost as versatile as hydrogen; and as a noble gas, it’s the least reactive. Helium offers a number of advantages over nitrogen, without needing hydrogen’s risk management or special equipment. Its most significant disadvantage is cost: helium is fairly expensive due to its high demand.

Important!  A note on helium as a resource, as there has been some misplaced concern about “depleting” the planet’s helium supply – it’s not possible.
As a product of abundant underground radioactive alpha decay, helium accumulates in pockets which replenish themselves (unlike other gases). It then escapes the planet’s gravity at a constant rate due to its low mass, so the earth will continue to produce and lose helium at about the same rate for millions of years whether we use it or not.
That means the supply of helium is not a depleting well whose bottom draws ever nearer. The most accurate analogy would be a flowing river from which we can draw a steady, but finite amount.

Flow Rate

Your flow rate can be controlled in two modes: constant flow or constant pressure. We recommend constant flow, because constant pressure leads to wide variations in flow, and most of calculations in GC are based on flow rate. Each carrier gas can operate within a specific range of velocities without losing column efficiency.

Column efficiency is a measure of how much benefit you’re getting out of the column. Think of it like this – if you blast your carrier gas through the column, the molecules will be travelling too quickly to interact with the column. Those interactions are responsible for separation, so your peaks will be closer together than you’d expect from just shortening the run.

Nitrogen gas has the best column efficiency between 10 and 15 cm/s, reaching a maximum at about 12 cm/s. Helium’s optimum column efficiency ranges from 15 to 40 cm/s, while peaking at 20 cm/s. Hydrogen has the largest optimum range, spanning 20 to 70 cm/s, with best results at about 40cm/s.

GC Column

Choosing a column might seem complicated, but it’s really no different than the experience you might have when buying toothpaste. There seems to be a staggering number of choices! But once you cut past all of the fluff, you discover that they’re all just slight variations on two or three products made by the same company. So we’re going to make the decision as straightforward as possible, because if you wanted information overload, you’d probably be reading their product sheets instead.

First, let’s choose a stationary phase by determining whether your samples are normally polar or nonpolar.

  • Nonpolar: Use a polydimethylsiloxane (PDMS) column
  • Polar: Use a polyethylene glycol (PEG) column
  • Somewhere in between: Use a polyethylene glycol (PEG) column
  • Combination / unknown: Use a 50% phenyl column
  • Quant only, no separation: Choose the opposite of your sample (faster analysis)

Next, we can turn our attention to the film thickness. If your sample identity and concentration are usually unknown to you, choose a thicker film (about 1 µm). You’ll gain better separation and a greater tolerance for high concentration samples at a slight cost in speed. If you know your analytes and already have good separation, shorten your run times with a thinner film (about 0.1 to 0.25 µm).

A 0.25 mm inner diameter is ideal for most labs. Deviating from this without good reason can introduce problems.

A standard column length is 30 meters. A longer column will give you better separation, but slow your analysis times. If your configuration already has a good separation efficiency, reduce your run times by shortening your column. If you’re not familiar with column cutting, ThermoFisher has an excellent guide.


As the GC’s temperature rises, more components of the sample will enter the gas phase and travel down the column. If you’re not getting enough separation, a temperature ramp can introduce sample components more gradually. Ramps can also be used to sharpen peaks for better quantification. But keep in mind that molecules can become unstable and break apart if the temperature is too high. That also includes your column coatings, so always check its specifications before raising your oven temperature.

Raising the temperature beyond your sample’s thermal stability will cause it to degrade, and molecules that are too large to enter the gas phase directly will only appear as fragments. It’s important to recognize this when it occurs to prevent making false identifications. Although thermal degradation will make any peak integration meaningless,  with enough experience you’ll be able to identify the parent molecule from the puzzle pieces. The next technique – derivatization – is especially useful in these situations.

Derivatization / Internal Standardization

Sometimes you might encounter a sample that seems like it could almost run, but doesn’t. It may go undetected, fragment as described above, or leave molehills instead of mountains on your chromatograph. These aren’t issues that can be fixed by making adjustments to your GC, and this is where derivatization comes in. It picks up where other methods leave off: so long as your sample doesn’t run the risk of out-right damage to the instrument, you have options to make it viable.

By subjecting the sample to a chemical reaction, you can change the chemical structure into a derivative that’s better suited for analysis. But derivatization isn’t just for samples you can’t run – you can also use it to produce derivatives with shorter analysis times! There are more reactions than we could ever list here – entire books have been filled with the possibilities. Fortunately for you, we have those books! Contact us for assistance if you need help with this technique.

Internal standardization is a technique that’s better suited to GC than GC-MS. This can help you to determine something elusive that can’t normally be detected using FID – like the precise concentration of water in a set of serial dilutions. By spiking your water with a detectable compound (one which won’t interfere with further analysis), you can quantify concentrations much more accurately.

Sample Concentration

For both data quality and instrument longevity, you should aim to use the smallest detectable sample size possible. This is usually within the 1-100 ppm range, favoring the lower end. Dilution calculations are very straight-forward: m1v1=m2v– just remember to account for the dilution factor in your final analysis.

Some labs won’t always have total control over the concentration of the samples they receive, or even prior knowledge of their contents. This is normally the case in forensic labs. In these cases, other methods can estimate the sample concentrations before introducing it to the sensitive components of the GC. If you place a highly concentrated sample on an FT-IR or a UV-Vis, the worst that can happen is a saturated graph.

Concluding Points

While your GC offers plenty of room for optimization, all of these factors interact with one another – so take care to avoid making multiple changes at once. For example: while changing your carrier gas and adjusting your flow rate are both ways you can improve your performance, making both of those changes together can actually worsen your performance.

This guide is intended to help you get the biggest improvement possible in the shortest amount of time. But the reality is that bringing your GC to highest potential will require a bigger time investment, some careful planning, and plenty of calculations. But that’s what we’re here for, so don’t hesitate to contact us. If you’re the type who likes to dive into the details, we can provide you with the resources and guidance you need. If not, we can schedule a visit and get your GC running better than ever.


Troubleshooting Gas Chromatography & GC-MS

My peaks keep showing up later and later every time I run a sample! (Retention Creep)

You probably need to change your septum. Certain brands wear out very fast, so this can catch you off-guard after switching if you’re used to a more durable brand. If changing the septum doesn’t fix the issue, secure or replace other vulnerable seals. If that still doesn’t fix it, ensure that your gas pressure and oven temperatures are where they should be. Also ensure that the run is being started consistently every time.

My peaks aren’t sharp enough to quantify! (Peak Tailing / Fronting)

As long as your installation hasn’t changed recently, this usually indicates a problem with the sample itself. Your sample concentration might be too high for that compound, or the sample could have poor compatibility with the column. Try diluting the sample, derivatization, or changing the column.

My peaks are running together! (Poor separation)

Follow the GC Optimization guide on this page to learn how you can improve your peak resolution and run time.

My chromatograms trail up into an S shape at the end of the run! (Column Bleed)

If you start to notice column bleed, it generally means that you’re falling behind on your maintenance, so it won’t hurt to give your GC a full tune-up. First, bake out your column by running it at the flow rate maximum and isothermal temperature limit for about 20 minutes. Start by replacing your short-lived components (seals, septa, and o-rings), then change your gas filters and trim your column ends by a few cm. Reassemble everything with extra care, and double check for leaks. If the bleed still continues, then your column is just worn out and needs replaced.

My peaks have peaks! (Split Peaks)

This is usually a problem with injection or vaporization. If it’s happening consistently for multiple users, then check ensure that the injector port is reaching the correct temperature, and verify that your solvent is compatible with your column. If it’s more intermittent, try injecting a lower volume more slowly (1 ppm & 1 µL should  be your guideline). If this still doesn’t fix it, consider adding wool to the injection port. This increases the surface area and aids rapid dispersion and vaporization.

My analyte isn’t showing up! (Unsuitable Samples)

GC-MS is a powerful technique, but it does have limitations. A general guideline is: if you can’t smell it, you can’t run it. Samples with high molecular weights (such as a heavy oils) are also unsuitable. When a molecule is too heavy to vaporize, you’ll never see the full molecule on the chromatograph. You’ll only see fragments of its thermal degradation – and you’ll continue seeing them until the sample fully degrades.

Important! Salts, ionic solutions, and metals are absolutely out of the question!

There’s something wrong with this GC Injection Syringe!

Even carefully maintained and properly used syringes wear out quickly with normal use, so it’s best to consider your syringes as consumables. Injection syringes are extremely fragile, and prone to failure at the slightest misuse. Mildly corrosive samples can jam up a syringe pump in a matter of seconds when introduced to the high temperatures of the injection port.

Proper cleaning includes flushing the syringe with an appropriate solvent after every sample. Follow this with water and several acetone rinses, and then place the syringe back into its case.

We recommend that you always have spares on-hand. Replacement parts can be helpful, but undetectable flaws can make it difficult to change them without accidental damage to the new parts. It doesn’t take many bent replacements before it becomes cheaper and less frustrating to simply have an extra.


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Autoclaves – Troubleshooting & DIY

Troubleshooting your Autoclaves

Important! This page is under active construction – please check back in a few days. Important!



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Micropipettes – Troubleshooting & DIY

Proper Care & Handling of Your Micropipettes

It’s hard to think of an instrument that’s more symbolic of modern science than micropipettes. They’re inexpensive, easy to use, and found in nearly every lab. But despite their apparent simplicity, most of their sources of error and malfunction aren’t very obvious. This page is intended as a quick reference of the most useful and esoteric information without making you waste your time skimming through non-essential information. Links to additional material are provided for anyone inclined to further reading.

Pipetting Dos and Don’ts

Do  Store the pipette vertically or at a downward angle in its intended holder.
Do  Always use a tip; pre-wet the tip; and change the tip before aspirating a different liquid.
Do  Clean the pipette exterior daily, and as required.
Do  Maintain stable temperature and humidity in your lab.
Do  Allow samples to equilibrate to room temperature before pipetting (when possible).

Don't  Lay the pipette down on the lab bench between uses.
Don't  Use the wrong size tip.
Don't  Turn the dial beyond the volume limits.
Don't  Drop the pipette.
Don't  Hold the pipette while it’s not in use.
Don't  Insert swabs into the pipette for cleaning.

Pipetting Technique

Pipetting is a skill that takes practice to master, and even pipette pros can benefit from an occasional review of the basics. Do you have flawless technique? Consider taking a moment to watch this excellent three minute video by Eppendorf to find out.


Troubleshooting your Micropipette

My pipette isn’t dispensing the right volumes consistently! (Poor accuracy/precision)

Errors, Contamination, & Overdue Maintenance
First, rule out user error by ensuring that everyone in the lab is pipetting with the same technique consistently. Next, consider the possibility of internal contamination. There will usually be visible clues – taking a dry, clean, lint-free white swab to the inside of the tip holder can be helpful. After this, consider the potential for internal wear. How old are the O-rings, and are they regularly exposed to organic solvents? How long has it been since your piston was lubricated with silicone grease? Your pipette may be overdue for basic maintenance.

Samples Requiring Special Techniques
Some samples may require you to use a special technique called reverse pipetting. These include:
Don't  Liquids with a tendency to foam.
Don't  Liquids with high viscosity.
Don't  Volumes of liquid near your pipette’s smallest deliverable amount.

Normal & Reverse Pipetting

How to Reverse Pipette
When pipetting normally, you press the plunger to the first stop, draw up your sample by releasing the plunger slowly, and then dispense the sample by pressing to the second stop to expel any lingering contents. In reverse pipetting, you begin by pressing the plunger all the way down to the second stop, and then draw up the sample by releasing the plunger slowly – completely. This places a reserve of liquid into the pipette tip that is greater than you intend to dispense. You then deliver the sample by only pressing the plunger to the first stop. The additional liquid will remain in the pipette tip. You can either return it to the original container or dispose of it into a waste receptacle as you see fit. Continue to hold the plunger at the first stop until you’re over the container you’ve chosen, and then press the plunger the rest of the way to the second stop before discarding the tip.

Off-Site Calibration
If you find that your pipette has good precision, but is delivering a consistently inaccurate volume, you will most likely need to start calibrating on-site. Most pipettes operate by air displacement, so a simple difference in location can bring your calibration out of spec. The density of the air in your lab is influenced by regional variations in air composition, combined with differences in elevation, humidity, and temperature. This can cause the air density at your lab to be very different from your calibration lab. Verdant only offers calibration on-site, because it’s the best practice.

When all else fails, blame the environment!
If you’ve ruled out contamination, your maintenance is up to date, your calibration is valid, everyone is consistently using the proper technique, then it may be your lab itself. If your lab’s environmental conditions are unstable from day to day, then your pipettes may be the first place you will see it. Micropipettes are high precision pneumatic instruments, which makes them especially vulnerable to lapses in environmental stability.

Contamination & Spills

Gradual contamination from proper use can be minimized by using filter tips. The negative pressures from pipetting will cause a small fraction of most samples to vaporize. Over time, this can lead to internal contamination of the pipette equivalent to a single spill. The filter tips form a line of defense by giving the sample somewhere else to deposit before reaching the internal components of your pipette. Evaluate the chemical properties of your samples to determine whether it’s prone to vaporize and deposit. If it is, you should pipette your samples with filter tips. If you’re not sure, you can always ask us.

Spills include any direct accidental contamination, such as pipetting with no tip, or laying the pipette on its side with a sample in the tip. If this happens, the pipette must be decontaminated, which will include checking and possibly repeating its calibration.

Important! The Forbidden Texts – Not for use in GLP/GMP labs Important!

Many pipettes are discarded after their calibration expires, because it’s often cheaper to replace than to re-calibrate. At Verdant, we strive to keep these out of landfills by bundling pipette coverage with other services at the same cost.

Outside of GLP/GMP settings, users can service their own pipettes as they see fit. But the absence of regulation doesn’t diminish the importance of a pipette’s accuracy and reproducibility. That’s why we’ve chosen to provide complete instructions on how to disassemble, clean, and calibrate a pipette.

Important! Disassembly procedures and calibration adjustment location vary from one manufacturer to another. It would be impossible to explain them all, so instruction manuals for most common pipette models are provided at the end of this section.

Decontaminating a pipette

Before servicing a pipette, it will need to be decontaminated, which will require that it be disassembled. And once a pipette has been taken apart, its calibration will need verified – and possibly repeated – before it’s put back into service. So there are three steps to a decontamination – cleaning, maintenance, and calibration. These steps should be performed every three months, or any time a pipette is either known or suspected to be contaminated. A full calibration (not just a check) should be performed annually.

Pipettes used for radioactive samples
Check the pipette for radioactivity using a calibrated Geiger counter. The standard maximum acceptable limits are 10 Bq for ɣ-radiation (gamma), 10 Bq for ß-radiation (beta), and 25 Bq for X-ray radiation. If the pipette exceeds these levels, it must be disposed of as radioactive waste in accordance with the above laws and policies. Otherwise, you may continue with the decontamination. There are no special procedures to reduce radioactivity. Cleaning a pipette contaminated by radioactive material only involves dilution and physical removal, by wiping the radioactive material away with a strong detergent and distilled water.

Important! If the institutional policies and laws governing your laboratory or physical location set a different maximum limit on radioactivity, always follow the lower (safer) limit.

Purchase or prepare a cleaning solution appropriate for your samples. This should be a mild industrial detergent which – depending on your sample – may include an oxidizing agent, a disinfectant, or an enzymatic cleaner. For pipettes which only deliver organic solvents, isopropyl alcohol followed by a 60 °C evaporation may be adequate.

1) First, soak lint-free tissues (such as Kimwipes) in the cleaning solution, and use them to wipe the body of the pipette.
2) Next, spray the disassembled components with the cleaning solution.
3) Dip a sponge-tip or lint-free swab in the cleaning solution, and then clean the inside of the tip-holder (the component to which the pipette tips attach).
4) Clean any O-rings and piston seals before discarding them into an appropriate receptacle for contaminated waste. These are to be replaced during reassembly.
5) Spray, wipe, or swab all remaining parts with the cleaning solution.
6) Repeat the above cleaning process for all components with distilled water (rinsing thoroughly where possible).
7) Follow this with isopropyl alcohol.
8) Leave the pipettes disassembled to dry in an oven set to 60 °C, or in a fume hood purged by high flow dry nitrogen for one hour. Otherwise, leave the pipettes covered by lint-free tissue, in a dust-free environment under positive pressure (such as a fume hood) if possible.

Important! When cleaning multiple pipettes at the same time, ensure that the parts are kept together with the original pipette. If you do swap parts or lose track, the pipette will need a complete re-calibration – not just a check.
Important! A sonicator bath can replace any physical cleaning steps above, saving yourself some labor and disposable cleaning items.
Important! Planning ahead can make your life easier: protein, DNA, or RNase samples are easiest to clean with an autoclave. If an autoclave safe pipette isn’t available, a mild detergent with an oxidizer such as hydrogen peroxide will suffice.
Important! Do not use alcohols or aprotic solvents. These will precipitate and this will complicate their removal.
Important! Do not expose your pipette to UV light to remove DNA. This will quickly degrade all types of plastic and shorten the life of your pipette.

Calibrating a Micropipette in Accordance with ISO 8655

Your pipette should be calibrated every twelve months, and checked every three months (called an as-found calibration). In the interest of keeping this article at a manageable length, please refer to this pamphlet by Gilson. It is the most well-written literature available: thoroughly explaining current best practices that meet or exceed ISO 8655.

Disassembly, Reassembly, and Calibration Specifics by Model

Instructions for the disassembly, reassembly, and calibration vary from one model to another. Instruction manuals are provided below for the most common pipette models.

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Analytical Balances – Troubleshooting & DIY

Troubleshooting Your Balance

The first step of troubleshooting is to clean your balance. To do this, you will need rubber gloves, fine tweezers, an anti-static bristle brush, canned air, Kimwipes, and any anhydrous alcohol. Once you’ve assembled your supplies, unplug the device and remove any batteries.

If any liquid is spilled, soak up the bulk with Kimwipes and then carefully remove the rest with lint free swabs.

Next, remove the platform and carefully sweep away any dust and debris using the anti-static brush. Use canned air to blow out any surfaces that are hard to reach. Do not pick the balance up and shake it, the internal components are extremely fragile.

Important! In place of an anti-static brush, you can use any bristle brush in combination with an anti-static gun, so long as the device is unpowered.

After any solid debris has been removed, wet a Kimwipe with the anhydrous alcohol and gently wipe the exposed surfaces of the balance. Gently use a cotton swab dipped in alcohol to clean deep surfaces and grooves. Allow adequate time for the alcohol to fully dry before replacing the weighing platform or your measurements will be impacted by a slow drift until drying is complete.

Important! Methanol, ethanol, and isopropanol are all suitable as long as they don’t contain any percentage of water. Do not use acetone or you will mar the plastic casing.

Once the device is clean, you can begin to check for common problems.

In some older devices, calibration weights are maneuvered into position by sliding a lever. If this lever is bumped accidentally, it can lead to inaccurate measurements. The process is automatic in most newer balances, but you may need to reduce the frequency as certain models auto-calibrate themselves every 15 minutes by default.

Interfacing With Your Balance

Interfacing with a balance is one of the most powerful ways to boost productivity. Interfacing is a way of letting your computer communicate with your balance, which means that your computer can tell your balance what to do, and request information. Anything you can do with a balance can be controlled by a computer, and modern analytical balances ship with this ability built-in.

You can read the balance output, tare, clear, print, calibrate, alter the settings, and even move the draft shield if it’s mechanized – all from your computer. You can even do it from home! This empowers you to automate repetitive tasks, and monitor the change in mass over extended periods of time (∂m/∂t) without expensive special equipment.

Most balances have separate manuals specifically for interfacing. Although these aren’t normally packaged with the balance, they’re usually available online or by request. Unfortunately, most instrument manufacturers have a frustrating tendency to require cords with awkwardly shaped connectors, so your first hurdle will be to acquire that cable. These can be extremely expensive from the manufacturer. For example, the “MiniMettler” is priced at approximately $350. We recommend searching for an aftermarket knock-off.

Once your computer is connected, you’ll need to choose software to communicate with the balance. The manufacturer’s software is a good starting point. These can usually perform basic tasks, but lack any useful customization. Most scientists will need the flexibility offered by a Hyperterminal client such as Realterm. These allow the scientist to communicate directly with the balance, and can automatically run simple scripts at regular intervals.

The most powerful option with the greatest flexibility would be data acquisition software, such as LabVIEW or DASYLab. These allow you to fully customize the collection and analysis of data without actually needing to program. The price range for a full commercial license can exceed $1000, but non-commercial licenses are usually available at a reduced price for academic & non-profit labs.


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DIY & Self-Service

Do-It-Yourself (DIY) & Self-Service Page





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