Translator for HPLC HINTS and TIPS for Chromatographers

Saturday, December 20, 2014

HPLC Column PORE SIZE (or Pore Diameter) and Retention Time

Think of your typical porous bare silica support as a big sponge. All of those holes are where the sample will migrate through before emerging out the other side. The size and number of those holes relate to retention time. Besides particle size (diameter), pore size is one of the most important characteristics of silica based chromatography supports.


The pore size or pore diameter (often expressed in Angstroms) is inversely related to the surface area of the support. The larger the surface area of the support (smaller pore size), the longer the retention of the sample. For small drug molecule samples under 1,000 daltons we often use high surface area supports with pore sizes between 60 and 120 Angstroms (~ 500 to 300 square meters per gram). These provide high retention characteristics useful in separating apart many compounds in one run. For larger molecules we employ supports with larger pore sizes (~300 Angstroms). Note: Pore size is often determined using the BET Nitrogen adsorption/desorption equation. Due to endcapping of the support (e.g. C8 or C18), the actual value obtained is often 20-30% less than the original value.

When comparing bare silica columns or trying to identify similar columns for use in a method, pore size must be considered. Manufacturer's publish the pore size in Angstroms (*sometimes in nm) for their different supports. Choosing columns with similar pore sizes will often provide similar retention characteristics. Accurate column void volumes are needed to calculate many standard chromatography performance calculations, but they are also critical for determining if you have any retention at all. T zero and the (k') K prime, capacity or retention factors, all rely on accurate column void volume values. It is one of the first calculations you must make BEFORE developing or running any HPLC method [link to more info: http://hplctips.blogspot.com/2011/05/determination-of-hplc-column-dead-time.html]. Once determined, you must follow up by injecting an unretained sample into the column to determine the actual value.

Saturday, November 15, 2014

Syringe Filter Selection for HPLC or LC/MS samples



This article will address the use of disposable female, Luer-compatible, syringe filters without built-in pre-filters for the filtration of individual samples into vials for HPLC or LC/MS analysis. - Note: 96 or 384 multi-well filtering plates provide for a better solution when large quantities of samples need to be filtered.


The choice of syringe filter depends on the: filter size (volume) of your sample, the chemical compatibility of the housing and membrane and desired pore size. Selection of the wrong filter size can result in too much sample holdup volume (loss of sample on filter) or overloading of the filter (allowing unfiltered material to pass through). If a membrane or housing is chosen which is not chemically compatible with your solution, then contamination of the sample or rupture of the assembly can result. Choosing a filter with too large a pore size can result in material passing through it which could clog or contaminate the solution (i.e. plug an HPLC system or result in a loss of sterility of a solution). Protein binding affinity is another characteristic of filter membranes and if you are filtering samples of biological interest, then you will also want to consider this specification in your selection criteria too (though it will not be discussed in this article).



Syringe Filter Size:


Filters are available in a variety of sizes which are generally in a disc shape and described by their diameter. Common sizes available for chromatography samples include: 3mm, 13mm and 25mm (~25-30mm) diameter discs. The larger the diameter of the disc, the larger the sample capacity, cross sectional surface area and potential hold-up volume of the sample on the filter. 


Hold-up volume is important because some of the sample will be retained inside the membrane and/or filter housing. If too large a filter is selected, samples with small volumes could be lost entirely in the hold-up volume on the membrane. Smaller filters have lower hold-up volumes. To extract as much sample as possible, be sure and use a post-filtration air purge to reduce the total hold-up volume.


If the volume of the sample you wish to filter is under 1ml, then a 3mm filter may provide the lowest hold-up volume and require the smallest amount of solution. To filter samples between 1ml and 10ml, the 13mm diameter filter provides a good balance between hold-up volume and large filter surface area. Larger sample volumes from 5ml to 50ml are often filtered through the more common 25mm diameter filters (~4 times the filtration area as a 13mm disc).




Chemical Compatibility:


Membrane Material: This is where you really must consult the manufacturer’s own documentation for the most compatible filter membrane for both your sample and the solution that will flow through the filter. To simplify the selection criteria, we can make some generalizations about some of the different types available:


Cellulose Acetate (CA): Use with aqueous solutions and a few hydrocarbons only. Low protein binding so good for many biological samples. Not compatible with ACN or DMSO. Can be autoclaved.


Nylon: Great general purpose material and compatible with many HPLC solvents (including THF, alcohols, ACN), but not strong acids. Nylon has a high affinity to bind proteins. Can be autoclaved.


Polysulfone / Polyethersulfone Variants (PS / PES): Commonly used with tissue culture and ion chromatography samples. Stable with many strong bases and alcohols, but few HPLC solvents (as it is hydrophilic). Low backpressure and low protein binding. Not compatible with ACN. Can be autoclaved.


Polypropylene (PP): General purpose hydrophilic material with resistance to most acids, bases, DMF, DMSO and alcohols. Not recommended for use with hydrocarbons, esters or solvents such as ACN. Can be autoclaved.


Polyvinylidene difluoride (PVDF): Hydrophilic material with broad compatibility. Often a good choice for use with alcohols, hydrocarbons, biomolecules, ether and ACN. Low protein binding. Can be autoclaved.


Polytetrafluoroethylene (PTFE): Reported in most brochures to be chemically resistant to almost all solvents, strong acids and bases. Hydrophobic membrane should be pre-wetted when used with aqueous solutions. Low protein binding and very strong. Can be autoclaved.

Most chromatography grade syringe filters are constructed of either HDPE or PP. These materials are compatible with a wide range of HPLC solvents and both offer low levels of extractables. HDPE has been reported to be more chemically compatible with aqueous basic solutions of NH4OH than PP.

Pore Size:


This will depend on your application and a number of different pore sizes are commonly available from vendors (1 micron, 0.8, 0.45 and 0.22 micron are the most common): 

For example, is sterilization of the fluid the goal? If so, a 0.22 micron filter is generally accepted as the best choice.  


For most chromatography or LC-MS applications either a 0.45 or 0.22 micron filters are preferred.




Summary:


  • Please refer to the various manufacturers data sheets to select an appropriate syringe filter with: (1) a low hold-up volume; (2) large enough size for the volume of sample; (3) which is chemically compatible with the solution and material you are going to inject through it and (4) lowest protein binding affinity (if applicable).
     
  • To reduce the hold-up volume, use a post-filtration air purge to empty the filter.
     
  • Minimize contamination from extractables (in the plastic) by pre-rinsing the filter membrane with some of the clean solution. This can reduce the amount of detectable extractables in your sample. PTFE based membranes have some of the lowest extractable levels so consider their use if this is an issue.
     
  • If analyte binding is a concern, select one of the membranes which has the lowest binding affinity such as PVDF or PTFE.

Saturday, October 11, 2014

Appropriate Mixer Volume for HPLC and UHPLC Applications

For gradient analysis, most HPLC (UHPLC) systems incorporate a solvent mixer which is designed to balance the requirements of moderate dwell volume, low noise and mixing efficiency. Depending on the method run, the ideal mixer's volume may in fact be completely different than the one installed in your chromatography system. Here are some very general guidelines to determine the appropriate mixer volume for your HPLC system. Note: Since many types of mixer designs exist, these are guidelines only.

HPLC System Mixer Volume Choices; Which Size Is Best?

SMALL: Fast or ultrahigh speed separations using low volume, small particle columns. These types of applications depend on a low dwell volume mixer. To achieve this, your HPLC system should be plumbed with narrow bore capillary tubing (example: 0.005" ID; 0.12mm ID) and include a gradient mixer with a volume of less than 100 ul (example: ~35 ul is rather common size). *Don't forget to address the dwell volume of the autosampler, loop and flow cell too when optimizing your system. 

LARGE: High Sensitivity Analysis: Gradient analysis where sensitivity is key, benefit from larger volume mixers to minimize contributions of any UV absorbing additives (e.g. TFA) and turbulence in the flow. Traditional 300 to 750 ul mixers often work well in these applications, provided that the column volumes are also large. Smaller column volumes will require smaller mixer volumes to reduce the added dwell effect.

MEDIUM: Routine HPLC Analysis: Typical analytical separations using 3 to 5 mm ID columns (x 100mm or longer) usually benefit from modest sized mixers within a range of 200 to 400 ul volume. For these applications, I often start with a recommendation to use a mixer which has 10% of the columns volume as a starting point. For a typical 4.6 x 250mm, 5 micron porous support column, which has about 3mls of volume, a 300 ul volume mixer should provide enough volume for routine gradient analysis. As mentioned before, the type of mixer, column volume, flow rate and mobile phase characteristics will help suggest the most applicable size for your application. Remember, these are general guidelines only. Your exact application must be taken into account to determine which size is best.

Saturday, September 6, 2014

Common Causes of Baseline Noise in HPLC, UHPLC.



Achieving a flat baseline which does not exhibit spikes, ghost peaks, drift or wander in an unpredictable manner should be a primary goal when performing HPLC analysis or developing methods. Methods which result in flat baselines and have well defined, sharp peaks allow for accurate sample area integration. Integration algorithms perform poorly in quantifying peaks on sloped, drifting or noisy baselines. Excessive baseline noise contributes to many problems, including poor quantitation, high %RSD errors, peak identification errors, retention time variation and many other critical problems. Properly developed HPLC methods are reproducible methods which apply and utilize good chromatography fundamentals. 


Note: A lack of proper training in the operation of the HPLC system, improper start-up or poor quality maintenance of the chromatograph (Examples: failure to degas and purge the system lines before use; an air bubble stuck in a check valve, a bad detector lamp or a leak will often result in baseline noise) are the main causes of noise. Your HPLC system must be optimized for your specific application. Be sure and allow time for the mobile phase to reach full equilibration with the system before starting any analysis.

In this article, we will discuss how temperature fluctuations, inadequate mixing, inadequate degassing and flow cell contamination can result in excessive baseline noise. We will provide suggestions on how to reduce or eliminate these problems.
TEMPERATURE FLUCTUATIONS:
To obtain reproducible results, the temperature of the hplc column must be kept constant during each analysis. Laboratory room temperatures often vary by several degrees during the course of one day and these changes will often change the retention characteristics of the sample(s). The 'On' and 'Off' cycling of power from an air conditioner or heating unit will often cause the baseline to drift in a cyclical manner, up and down, during the day (this can often be seen as a clear sine wave pattern when you zoom-in to study the baseline trace over time). Temperature also changes the refractive index of the mobile phase. Light based detectors (UV/VIS, RI...) will show this change as drift up or down). In some cases, a temperature change of plus or minus one degree C from run-to-run can cause changes in retention times which effect reliability of the method. 

To reduce temperature fluctuations, you must control the temperature of the column and mobile phase (if applicable) during the analysis. This is most commonly done by: (a) using equilibrated mobile phase at the start of the day or analysis, (b) keeping the interconnecting lines as short as possible (esp. any which exit the column and go to detectors/flow cells), (c) insulating any stainless steel lines with plastic tubing to reduce heat loss and (d) using a thermostatted column compartment to maintain the column at a single set temperature throughout the day. Control of the column temperature will remove 'temperature' as a variable from your analysis. Temperature should be a constant run to run, not a variable. Be sure and document the temperature selected as part of your method.
INADEQUATE MOBILE PHASE MIXING:
Both high pressure (with separate pumps) and low pressure pumping (one pump with a proportioning valve module) systems depend on efficient mixing to reduce noise. For gradient analysis, failure to completely mix the mobile phase solution before it enters the HPLC column often results in excessive baseline noise, spikes and poor reproducibility. "Mixing" is often accomplished directly in a mixer installed in the flow path of an HPLC pump. The associated noise and ripple of incomplete mixing can reduce the limit of detection (LOD) and increase integration error. This mixer is often a static mixer (a simple 'Tee', a tube filled with baffles, a frit or beads, valve orifice or microfluidic device) of low volume design for chromatography use, but allows adequate mixing of the liquids within a prescribed flow rate range. The best mixers incorporate longitudinal and radial mixing in-line. A mixer with too low a volume or of insufficient design can result in poor mixing of the mobile phase (note: incorrect solvent compressibility settings can also cause mixing and noise problems too). To reduce mixing problems, first insure that the mobile phases used are fully soluble with each other. Next, make sure that any mixer used is appropriate for the flow rates and volumes you will be using. Monitor the baseline for drift, ripple and artifacts in real time to spot problems and make adjustments to correct them. 
INADEQUATE MOBILE PHASE DEGASSING:
For the best results, continuously degas your mobile phase. Reducing the amount of gas will also improve signal to noise levels of detection, reduce drift and reduce pump cavitation. If you are using an electronic vacuum degassing module, make sure it is maintained and working 100%. A faulty degasser may cause more damage (contamination) to your system and methods. Maintain and Repair them just as you do for your other instrument modules. Gas bubbles may cause check valves to malfunction (get stuck), baseline noise spikes to appear randomly, flow rates and/or pressures to become irregular, detector outputs to show high levels of noise (from air in the flow cell) and also cause the loss of prime or cavitation in pumps. To achieve the best balance of low noise levels and high reliability, both aqueous and organic mobile phases should be fully degassed before and during use. This can be accomplished through stand-alone inline vacuum degassing modules or through gentle continuous helium gas sparging (*Helium makes an excellent choice of gas as it is not soluble in the mobile phase. Never use Nitrogen or Argon gas, they are soluble in the liquid!). In all cases, degassing must be continuous (not just done one time). Continuous degassing reduces cyclical noise and signal variations. For this reason, I do not recommend using ultrasonic baths to degass mobile phase solutions as these are not used in a continuous mode. The mobile phase solution starts to re-absorb gas as soon as you stop sonicating the solution. This results in continuous baseline drift (up and down).
Removal of gasses is critical to the function of a modern HPLC pumping system. The liquids used are compressed to very high levels which forces out solubilized gas from the solutions. This is best accomplished before the liquid is transferred into the pump. These gas bubbles must be minimized to achieve desirable baselines. *Even if you use a high pressure pumping system, an inline degassing system reduces the amount of noise and baseline drift. Properly maintain and service your degasser to insure compliant operation. IOW: Whichever method you use, always degas your mobile phase solutions.
FLOW CELLS:
One other less common cause of baseline spikes and random noise is due to either a dirty flow cell (i.e. the windows) or an air bubble trapped inside the flow cell. If the flow cell is suspected of having one of these problems, then it should be carefully rinsed or flushed out with an appropriate mixture of suitable solutions to expel the air bubble or remove the contamination. If possible, keep a spare, 'known good' flow cell on hand to swap out for troubleshooting purposes. This can help to quickly determine where the problem is. This flow cell must be the exact same size and type (volume and path length) for this purpose. If the cell's windows are contaminated and flushing does not restore them, then many manufacturer's offer kits which allow you to replace the windows and gaskets used. Warning: When attempting to clean or repair any flow cell, be sure and work within the manufacturer's operational specifications for the specific flow cell. Some flow cells are not designed to withstand even very low back pressure and damage can result if you exceed their maximum pressure or chemical rating.

Many other types of problems not mentioned in this short article can also cause baseline noise. For example, a sticking inlet or outlet valve on the pump, worn out detector lamp(s) or detector electrode (EC) can induce noise. In all cases, the cause must be investigated in a logical, step-wise manner. Demonstrate what is working and rule out items one-by-one.

Reference: http://hplctips.blogspot.com/2014/01/diagnosing-troubleshooting-hplc.html

Saturday, July 12, 2014

USP Guideline Note: HPLC Column Diameter Changes to Maintain Linear Velocity


USP Allowed Variations in HPLC Column Diameter (*USP 32, Second Supplement, Dec 1, 2009). In the previous USP version, a change of up to 50% of the flow rate was allowed. This has been changed in the more recent version. A wide range of column diameter changes are now allowed, provided that the linear velocity is kept constant. *We addressed the effect of changing column diameter on flow rate in a previous blog post, but this time I have also expanded on the calculation by including the extra variable for column length (L1 and L2) as well.




Linear Velocity Formula:

   New Linear Flow Rate2 = Flow Rate1 x (L2 x D22) / (L1 x D12)

Flow Rates are in ml/min.
L1 = Column Length (original) in mm.
L2 = Column Length (proposed) in mm.
D1 = Column Diameter (original) in mm.
D2 = Column Diameter (proposed) in mm.

Example #1:
Original column is a 150mm x 4.6mm (L x ID) used at 1.000 ml min. We would like to determine what the equivalent flow rate (F2) would be for a column which is 150mm x 2.1mm (L x ID) to maintain the same linear velocity. This is a proposed change in column diameter of > 50% so it would not have been allowed under the old guidelines. The newer guidelines take into account that with the same particle size, changing the column diameter will not change the chromatography if the linear velocity is maintained as before. Let’s calculate the new flow rate using the formula above.
1.000 x (150 x 2.12) / (150 x 4.62) = F2
                    1 x (661.50 / 3,174) = F2
                              0.208 ml/min = F2