DATA INTEGRITY AND ALCOA +

 

                        DATA INTEGRITY AND ALCOA + 

                        

Your  question ---

Q: I am familiar with the term ALCOA as it relates to data integrity, but lately, I have heard people refer to ALCOA+. Can you explain what impact this new acronym has on my company’s data integrity program?

ANSWER : -  Before we get into the reason for the additions to ALCOA, referred to as ALCOA+, we should review what both of the acronyms mean. The acronym ALCOA requires data be attributable, legible, contemporaneous, original, and accurate. The acronym ALCOA+ adds the concepts that, in addition to ALCOA, data also needs to be complete, consistent, enduring, and available. It is important to understand what each element of ALCOA and ALCOA+ mean in order to apply the concepts appropriately with respect to a company’s records. The following are some general definitions, paraphrased from the Pharmaceutical Inspection Co-operation Scheme (PIC/S) (1), that can be used for understanding the elements of ALCOA and ALCOA+:

• Attributable: The data generated or collected must be traceable back to the individual who generated the information.

• Legible: The data recorded must be readable and permanent.

• Contemporaneous: The results, measurements, etc. must be recorded at the time the work is performed.

• Original: Original or source data are the record, report, notebook etc. where the data point was initially recorded.

• Accurate: The data recorded must be complete, consistent, truthful, and representative of facts.

• Complete: Information that is critical to recreating and understanding an event. This would include any repeat or reanalysis performed on a laboratory test sample.

• Consistent: The data are presented, recorded, dated, or time-stamped in the expected and defined sequence.

• Enduring: The data or information must be maintained, intact, and accessible throughout their defined retention period.

• Available: The data or information must be able to be accessed at any time during the defined retention period.

Data integrity has always concerned regulatory authorities, but it is important to understand what is prompting the renewed discussion of ALCOA and the introduction of ALCOA+ when discussing data integrity issues. Many of the concepts for ALCOA have been captured in the regulations as far back as 1978. Since that time, the industry has changed dramatically. The generic-drug industry has grown and in the United States alone accounts for more than 80% of the prescriptions written today (2). Coupled with the emergence of biosimilars, virtual companies, contract manufacturing organizations, rapid advances in automation and information technology, and the globalization of the industry have resulted in reinterpretation of the attributes associated with maintaining the integrity of data throughout the product lifecycle, whether those data are generated from electronic, paper-based, or hybrid systems. In addition, there has been an increase in citations internationally by the FDA, European Medicines Agency (EMA), Medicines and Healthcare products Regulatory Agency (MHRA), World Health Organization (WHO), and other health authorities. These changes and increased violations have brought about a resurgence and need to reeducate the industry on the basic principles and concepts regarding the proper control of data used to support the quality and safety of medicines.

The ALCOA acronym has been used since the 1990s; however, the requirements governing data elements have been in regulations for a much longer period of time. EudraLex chapter 4 states, “Suitable controls should be implemented to ensure the accuracy, integrity, availability, and legibility of documents. Instruction documents should be free from errors and available in writing” (3). The US Code of Federal Regulations (CFR) refers to these elements in various sections of the regulations (4–8). An example for language speaking to the element of attributable is in 21 CFR 211.194(a)(7), which states, “The initials or signature of the person who performs each test and the date(s) the tests were performed.” Language in 21 CFR 58.130 (e) addresses the elements of contemporaneous and legible by stating, “All data generated during the conduct of a nonclinical laboratory study, except those that are generated by automated data collection systems, shall be recorded directly, promptly, and legibly in ink. All data entries shall be dated on the date of entry and signed or initialed by the person entering the data.”

Recent documents issued by WHO (9) and the PIC/S (1) have added to the original ALCOA attributes as indicated above. The PIC/S document actually states, “Some key concepts of GDocPs are summarized by the acronym ALCOA: Attributable, Legible, Contemporaneous, Original, and Accurate. The following attributes can be added to the list: Complete, Consistent, Enduring, and Available. Together, these expectations ensure that events are properly documented and the data can be used to support informed decisions.” WHO refers to ALCOA+ in the title of Appendix 1 to their 2018 document. The last two documents also address the concept of quality culture (10). The impact to your organization is that the quality culture must ensure that data supporting the quality and safety of your product must now meet the ALCOA+ elements in order to avoid regulatory citations for data integrity issues.

References

1. PIC/S, Draft Guidance: Good Practices for Data Management and Integrity in Regulated GMP/GDP Environments (PIC/S, August 2016).
2. statista.com 2019.
3. EC, EudraLex,The Rules Governing Medicinal Products in the European Union, Volume 4, Good Manufacturing Practice for Medicinal Products for Human and Veterinary Use, Chapter 4.
4.  21 CFR 11: Electronic Records; Electronic Signatures
5.  21 CFR 58: Good Laboratory Practice for Nonclinical Laboratory Studies
6.  21 CFR 211: Current Good Manufacturing Practice for Finished Pharmaceuticals
7.  21 CFR 212.50: Current Good Manufacturing for Positron Emission Tomography Drugs
8.  21 CFR 820: Quality System Regulation
9.  WHO, Annex 5, Guidance on Good Data and Record Management Practices (WHO, June 2016).
10. S. Schniepp, Pharmaceutical Technology 42 (10) 2018. 

NATURE OF A GOOD MOBILE PHASE

 

                            NATURE OF A GOOD MOBILE PHASE

ABOUT MOBILE PHASE

Mobile phase has been rightly termed as the lifeline of the HPLC system. It plays the important role of transport of the sample through the separation column and subsequently to the detector for identification of the separated components.

Mobile phase is seldom a single solvent. It consists of combination of water with organic solvents, aqueous buffers with polar solvents or mixtures of organic solvents in desired proportions. The two operational modes commonly used are:

Isocratic mode – in the Isocratic mode the composition of the mobile phase remains constant throughout the analytical run ,i.e, the proportion of solvents in the mixture is pre-decided and remains unchanged during analysis

Gradient Elution – the composition is varied by the software through the analytical run in a predetermined mode and at the end of the run the proportion of solvents is different from the initial proportion.

The objective of using different solvent mixtures is to achieve the desired polarity of mobile phase for complete miscibility of the sample and control interaction of sample components with the stationary phase to achieve the desired degree of resolution of separated component peaks in the shortest possible time.

In order to meet the required objectives the mobile phase should have the following essential features:

• The sample should be fully soluble in the mobile phase. Any insolubility will result in flow restrictions. Always check sample solubility in mobile phase before injection into the system.
• Mobile phase components should be non-hazardous and non-toxic. They should not pose any health hazard to the operator.
• Mobile phase shouId be inert towards sample constituents and the stationary phase. Any reactions can lead to formation of insoluble suspensions which can result in column blockages.
• The mobile phase should not give its own response on passing through the detector. In other words the detector signals should reflect only the response of the sample constituents. This, however, is not applicable in bulk property detectors such as refractive index detector which respond to overall changes in refractive index of the mobile phase containing the eluting compound.
• The mobile phase should be affordable and the proportion of solvents used should make the analysis economically viable.

It is important to mix the solvents of specified purity and from same source to get consistency of results and also to adopt the same off-line mixing technique to avoid errors due to heat of mixing errors. In case buffers are to be used always adjust the pH of the aqueous phase prior to making up the final volume with the organic phase.

HPLC technique holds great promise for applications in diverse fields. Control of operational conditions including consistency of mobile phase will ensure repeatability of results.

ROLE OF PH IN MOBILE PHASE CPMPLEAT GUIDLINES

                        ROLE OF PH IN MOBILE PHASE COMPLETE GUIDLINES

The mobile-phase pH can be a powerful tool to control retention and selectivity, but it can also get you in trouble if it is not controlled properly.

The adjustment of mobile-phase pH can be a powerful tool to obtain liquid chromatographic (LC) separations, but at the same time, poor control of the pH can be a source of serious problems with separations. In this installment of “LC Troubleshooting” I’d like to focus on mobile-phase pH and its role in retention and selectivity.

pH and Retention

In reversed-phase LC, retention is dominated by the overall hydrophobic or nonpolar nature of the analytes. Compounds that are more polar tend to have shorter retention times than their nonpolar counterparts. As a result, the elution order of the typical chromatogram proceeds from polar to nonpolar compounds. If all the sample components are neutral, the mobile-phase pH generally can be ignored as an important factor in retention. This situation is observed in the chromatograms of Figure 1, where the unlabeled peaks eluted between approximately 3 and 9 min are neutral compounds. Notice that the retention of the neutrals is not altered with a change in pH from 7.0 (Figure 1a) to 3.0 (Figure 1b).

Figure 1: Simulated chromatograms for a mixture of six neutral compounds (unlabeled peaks between approximately 3 and 9 min) and two carboxylic acids at (a) pH 7.0 and (b) pH 3.0. The peak at approximately 0.75 min is t0.

 

When ionizable analytes are present, dramatic changes in retention can take place when the mobile-phase pH is altered. In Figure 1a, the carboxylic acid components are strongly ionized at pH 7.0, so their negative charge makes them more polar than the same molecules at pH 3.0 (Figure 1b). Thus we can see that pH can have a significant affect on retention of ionizable compounds.

I often cite my “Nothing’s Magic” rule, which says that, in general, we can assume a continuous change in the appearance of a chromatogram between two observed conditions and often can safely extrapolate somewhat beyond the observation points. In the current example, this rule suggests that between pH 7.0 and 3.0, the retention times of the acids will fall between those of Figures 1a and 1b.

Retention of Acids and Bases

The changes in retention for acids and bases with a change in pH, not surprisingly, are in the opposite direction, as shown in Figure 2. In Figure 2a, we see that acids have good retention at low pH and poor retention at high pH, just as we observed in Figure 1. Just the opposite happens for bases (Figure 2b), because at high pH, bases are neutral and well retained, whereas at low pH they are ionized and poorly retained.

Figure 2: Generalized plots of retention versus mobile-phase pH for (a) acidic samples and (b) basic samples. These plots do not directly correspond to any of the example chromatograms, but rather show the general retention behavior of acids and bases as pH is changed.

 

 

At the midpoint of these curves is the pKa of the acid or base, where half of the molecules present are ionized and half are not ionized. Note that although both ionized and nonionized species are present at the pKa, only a single peak is observed. This occurs because the equilibrium between the two forms is so rapid compared to the time it takes for the sample to travel through the column that the analyte behaves chromatographically as the average of the molecules present. As the pH is shifted to a lower pH for acids, the portion of the total molecules present in the un-ionized form increases and the number of ionized molecules decreases, so the overall polarity of the acid decreases, resulting in longer retention times (moving left in Figure 2a). Bases have the opposite response to a change in pH.

Although the change in ionization is significant within approximately 1 pH unit of the pKa, you can see that the curves flatten out at both ends by the point the pH is 2 or more pH units above or below the pKa. It is generally considered that at 2 pH units from the pKa, an acid or base is fully ionized or ionization is fully suppressed. So the effect of a change in mobile-phase pH depends on the pH value compared to the pKa of a compound of interest. This means that if we want to use pH to adjust retention, this will be most effective within approximately 1­1.5 pH units of the pKa. On the other hand, for the most robust retention conditions, the mobile-phase pH should be >1.5 pH units from the compound’s pKa.

The Selectivity Power of pH

The real beauty of exploring mobile-phase pH during method development for samples that contain acids or bases is that the pH versus retention response curve varies somewhat from one compound to the next, especially if their pKa values differ. In practical terms, this means that significant changes in retention and peak spacing can be obtained with a change in mobile-phase pH. An example of this, as well as further elucidation of the generalized retention behavior of bases in Figure 2b, is shown in Figure 3 for separation of four substituted anilines (1). The peaks in Figure 3a are numbered and they are color-coded for all the peaks in Figure 3 for easier peak tracking. Analyte identities and pKa values for the peaks of Figure 3 are summarized in Table I. Note that the pKa values for these compounds vary from 2.66 to 3.98. The discussion above indicated that pH values within ±1.5 pH units of the pKa will be most effective at changing retention, so we’d expect to see the most peak movement in the pH range of (2.66–1.5) = 1.1 to (3.98 + 1.5) = 5.5. I have shown simulated chromatograms in Figure 3 of 2 ≤ pH ≤ 5.5; pH < 2 may cause column damage and pH > 5.5 showed almost no change from pH 5.5 for these analytes.

Figure 3: Simulated chromatograms for four substituted anilines at the pH values shown in the figure. Data of Table I of (reference 1), but adjusted for a 150 mm x 4.6 mm column size. Column type: StableBond CN column (Agilent); mobile phase: 25:75 buffer–methanol; buffer: 25 mM sodium citrate (pH ≥ 4.0) or potassium phosphate (pH < 4.0); flow rate: 1 mL/min; temperature: 35 °C. See Table I for compound identities and pKa values (2). The peak at ~1.7 min is t0.

 

Let’s track the movement of the various peaks and see how they are affected by pH. Peak 4 (black) with pKa = 2.66 will be ionized at pH 2.0, so it is least retained at this pH. As the pH is increased above pH 2.5, the nonionized species predominates and retention increases, as expected. By the point pH 4.0 is reached, ~1.5 pH units above the pKa, the ionization is suppressed (flat region to the right of Figure 2b), and little if any further change in retention is observed in the pH 4.0–5.5 chromatograms. Peak 3 (orange) has a similar pKa (2.75), so very similar behavior is seen for peaks 3 and 4, with stable retention above pH 4.0. Although the retention times of these two peaks change with pH, the selectivity, α, changes only a small amount (range of α = 1.36–1.40), so they nicely separated at all pH values shown. (Recall that α is the ratio of retention factors, k, for two adjacent peaks; in this case, α = k4/k3.)

We can similarly track peak 1 (green, pKa = 3.98) and peak 2 (red, pKa = 3.52). These peaks also move to higher retention times as the pH is increased and ionization subsequently is decreased. However, because the pKa values for peaks 1 and 2 are approximately 1 pH unit higher than those for peaks 3 and 4, the retention times do not stabilize until approximately 1 pH unit higher, as well, at pH 5.0 and above. Unfortunately, the selectivity between peaks 1 and 2 drops with increasing pH, from α = 1.64 at pH 2.0 to 1.05 at pH 5.5, so the separation of these peaks worsens with increased pH.

 

 

Mobile-Phase pH and Robustness

Many pharmaceutical laboratories are adopting the quality by design (QbD) guidelines (3) from the International Conference on Harmonization (ICH). These guidelines encourage establishing a design space that encompasses the boundaries of the process variable settings that provide acceptable quality. From a chromatographic standpoint, this means identifying the range that a chromatographic variable can be changed and will still provide acceptable analytical results. In the current context, this means performing robustness experiments to determine how much the mobile-phase pH can be varied and still provide adequate separation.

An example of a method with poor robustness to pH is shown in the chromatograms of Figure 4 for a sample of bile acids (4). In Figure 4a, all the peaks are separated to baseline at a mobile-phase pH of 5.1. However, when the pH is shifted to 5.2 (Figure 4b), the last two peaks merge into a poorly separated doublet (arrow). When a pH meter is used to measure the pH of a solution during pH adjustment, normal laboratory variation is typically ±0.05–0.1 pH units. When the pH of a buffer is adjusted by titration, differences such as those seen between Figures 4a and 4b would not be surprising when different batches of buffer were prepared. As a result, the separation would not be sufficiently robust if a mobile-phase pH value of 5.1 were specified.

Figure 4: Simulated separations of a sample of bile acids based on data of (reference 3). Mobile-phase pH: (a) 5.1, (b) 5.2.

 

Let’s consider how to establish satisfactory separation conditions for the sample of Figure 3 that have acceptable robustness. Generally, we like to have isocratic retention factors, k, of 2–10, but 1–20 usually is acceptable if 2–10 is not possible. Also, let’s specify that we want baseline resolution between peaks. In addition we would like a method that tolerates as much variation in pH as possible.

The separation at pH 2.0 in Figure 2a looks good, but the k-values for the first two peaks are <1, so there is a risk of interferences at the column dead time, t0, when real samples are run. At pH 2.5 (Figure 2b), k for the first peak is ~1, and plenty of separation is observed, so this pH is acceptable. At pH 3.0 (Figure 2c), peak 2 has moved partially past peak 3, and the separation is not adequate. At pH 2.75 (chromatogram not shown), peak 3 is just baseline separated after peak 2, so this is about the maximum acceptable pH; at all larger values of pH, peak merging is observed.

With acceptable separations seen over a range of 2.0 ≤ pH ≤ 2.75, we would probably set the default pH midway between these points-for example, at pH 2.35. During validation we could then run experiments at pH 2.05 and 2.65. If these conditions gave satisfactory separation (and other method performance requirements), we could define the method pH as 2.35 and allow adjustment of the pH by up to ±0.3 pH units. Such a statement in the method document would allow the analyst to adjust the pH by up to 0.3 pH units to achieve system suitability.

To ensure that the mobile phase is as stable as possible, we use a buffer in the mobile phase. Buffers are most effective ±1 pH unit from their pKa values. Phosphate has three pKa values, at 2.1, 7.2, and 12.3, so it would be an appropriate buffer to use in the pH 2.35 ± 0.3 range. Contrast this with acetate, with a pKa of 4.8, which would not be an appropriate buffer for the sample of Figure 3. On the other hand, if it were necessary to operate the method of Figure 4 at pH 5.1, acetate would be the right buffer to use, not phosphate. A buffer concentration of 20–30 mM is common for most LC methods, although concentrations as low as 5–10 mM are suitable with today’s high-purity silica columns.

Summary

Mobile-phase pH will have little effect on the retention of neutral compounds, but if ionizable compounds are present in a sample, pH control is necessary to stabilize retention. The pH of the mobile phase can be an extremely powerful tool to move peaks around in the chromatogram during method development, but the flip side is that it needs to be carefully controlled during routine analysis to maintain robust separation conditions. When selecting a buffer to control the mobile-phase pH, be sure to select one with a pKa value within 1 pH unit of the desired pH or it may not have enough buffering capacity to stabilize retention.

How does a C18 HPLC column work ?

    How does a C18 HPLC column work ?

Answer - A C18 column is a reverse phase HPLC column which has carbon chains bonded to the silica particles inside the column. The C18 tells you how long the carbon chains are, in this case there are 18 carbon atoms in each chain. This type of phase is hydrophobic and non polar molecules will interact with it when they pass though the column. So if you had a sample which had two components in and one was more hydrophobic than the other, the least hydrophobic component would elute first followed by the more hydrophobic molecule. To help drive these molecules off the column a mobile phase with a high solvent content is used so that the mobile phase is more hydrophobic than the column, this will cause the molecules to interact less with the column and stay in the mobile phase for longer, until they eventually elute out of the column and then travel to the detector for analysis.

CARRYOVER TROUBLE in HPLC and Action Steps

                        HPLC Carry over and its Solution to prevent

Carryover Discription

 Carryover is recognized as the presence of a small analyte peak that appears when a blank is injected following the injection of a sample that produces a large peak of the same analyte. When it occurs, peaks attributed to the previously analyzed sample may be observed in the subsequent chromatogram(s) which may co-elute or interfere with desired analytes. Shown below, the mass spectrum of a blank reveals the profile of a target analyte. It is one the most frustrating problems in HPLC.

Carryover usually amounts to a chemistry problem when coupled with certain sample injection flow path materials and hardware connections. What steps can we take to solve it?

Action Steps Overview

​

1. Classify the carryover
2. Replace the blank and vary the injection volume
3. Rule out the chromatography column
4. Check flow path fittings
5. Check autosampler rinse phase solvent(s)
6. Enable autosampler rinse mechanism
7. Change hardware
8. Addressing sample-specific carryover

Take Action

1. Classify the carryover

a. Classic or Constant

i. Classic Carryover – The observation is a regular reduction of carryover peak size as blanks are injected consecutively. For example, the first blank might have a peak that is 1% of the size of the original, and then the next drops by another factor of 100. This carryover is often caused by a mechanical area in the flow path (like the intersection between two surfaces in a valve head where tubing is inserted and the nut is swaged). Hold-up of some sample is progressively diluted upon injecting blanks, causing progressively less carryover with each blank run.

ii. Constant carryover – Not true carryover. A small peak is always present with all samples and blanks and does not diminish with each blank run. This is caused by some source of contamination. If the contaminant peak seems to increase or decrease with increasing and decreasing blank injection volumes, it is likely that the blank is contaminated. If not, the source of contamination is elsewhere in the system.

iii. Consider the importance of a ‘null-injection’ run. Shimadzu autosamplers offer the unique ability to perform a null-injection, which starts the chromatography run without injecting sample and without rotating the injection valve or high pressure valve of the autosampler. This simple test can help to determine whether the source of the problem is the injection event itself. A null run that produces no offending peak must indicate that the auto-sampler injection event is the source of the issue. It does not prove the source of the carryover within the autosampler, but it does narrow the problem down to the high pressure valve (HPV), its connections, and the outlet tubing connected to the column. Because Shimadzu autosamplers are needle-in-flow path design, this experiment rules out the sample loop and needle because eluent is always flowing through the loop and needle unless the valve is rotating or sample is being aspirated from a vial.

2. Replace the blank and vary the injection volume

a. It is important to rule out whether the blank itself may be contaminated. The easy way to check this is to prepare a fresh blank using a different matrix source (i.e. the solvent itself may be contaminated). In addition, varying the injection volume can be helpful –if you increase the injection volume of the blank and the peak area of the carryover peak increases as well, it is a strong indicator that the blank is contaminated. If you notice the size of the carryover peak remains constant, then most likely the carryover is located on the outside of the needle (see instructions for external autosampler rinsing). If you notice the size of the carryover peak decreasing with each subsequent injection then most likely the carryover is located inside of the needle and/or sample loop (see instructions for internal autosampler rinsing).

 

3. Rule out the chromatography column

a. The chromatography column may be the source of carryover, especially if it is fouling over time. The repeated injection of samples containing intractable materials or strongly retained constituents will slowly modify the characteristics of the column bonded phase with each injection. Gradient conditions may not effectively clean the column of such materials and the problem may worsen over time. The simple test for a problem column is to remove it from the system and replace it with a zero-dead-volume union. Do a sample run as normal, noting that there is no column to retain and differentiate sample constituents. Follow it with a blank run or two. Does carryover persist? If so, the problem is hardware related. If not, the problem lies with the chromatography column and you should replace it.

 

b. Another experiment can indicate whether the column is producing the carryover. Create a method that takes the pumps through a double gradient, as pictured below. The run begins with a sample injection. The gradient program moves from a low strength eluent condition to a high strength eluent condition, flushes the column at high strength, then returns to initial conditions for a couple of minutes. The same gradient is then repeated, but without a new injection occurring. In this way, the column bed is taken through a gradient of solvent strength to see if a carryover peak is again produced. If it is, it may indicate that the column packed bed or inlet frit are the source of carryover and a more aggressive and frequent column flush may be in order.

4. Check flow path fittings

a. This procedure generally is limited to the fittings on the HPV and all fittings downstream to the detector. Stainless steel fittings rarely are a problem, but PEEK fittings and tubing can slip under high pressures; for example, greater than approximately 4000 psi. You can check stainless steel fittings by slightly tightening them with a wrench. The best approach for PEEK fittings and tubing is to shut off the pump, loosen each fitting, push the tubing firmly to the bottom of the tube port, and re-tighten the fitting.

5. Check autosampler rinse phase solvent(s)

a. Purge the autosampler

i. This is a precautionary step to ensure all of the rinse lines leading supplying the autosampler are metering rinse solvent and not charged with air. It is possible that proper needle or seal rinsing is not occurring due to air in the lines.

ii. Use fresh rinse solvent - Replace the rinse phase solvents with a fresh batch and replace the rinse phase reservoirs with clean ones. After the rinse solvent is replaced, cycle the autosampler through the purge or rinse cycle several times to ensure that all traces of the previous rinse phase solvents have been removed.

b. Adjust rinse phase strength

i. A strong rinse phase is one that has an affinity for the sample constituents causing the carryover. For reversed phase chromatography where the organic solvent is the stronger, use more of it in the rinse mixture to ensure effective rinsing. Often it is effective to make the rinse phase chemistry roughly equivalent in strength to the mobile phase. Sometimes it is worth considering the use of 100% of the strong solvent (usually acetonitrile or ethanol). Isopropanol makes an excellent wash solvent for many applications (e.g. fatty acids) and proves to be more effective for removing contaminants than methanol or acetonitrile.

c. Adjust rinse pH

i. Consider that rinse phase pH is a form of chemistry ‘strength’. In some cases, analytes causing carryover may be quite sensitive to the pH of the rinse phase. For rinse mixtures, avoid using pH buffers or salts that leave a residue when evaporated. Never use phosphate buffers. Use volatile pH modifiers as one would use for LCMS mobile phases, like formic acid, acetic acid, ammonium formate, ammonium acetate, or ammonium hydroxide. Typically a 0.1–1% solution by volume is sufficient. Try adding acid or base to the wash solvent to see if the carryover is improved.

6. Enable autosampler rinse mechanism

a. Enable pre- and post-injection needle rinse. See the section at the end of this document for specific information about rinse types and rationale for use. Multi-rinse options allow for a combination of strong/weak rinse chemistries to be used for both external needle surface rinse and internal injection mechanism rinse.

 

7. Change hardware

a. Change needle, needle seal, injection loop, or HPV rotor

i. If you’re still trying to solve the carryover problem, it is highly likely that the problem is related to the autosampler hardware. At this point, you should substitute parts until you locate the problem. Often the easiest first step is to try a manual injection valve or to replace the autosampler with one that has not been exposed to the compound of interest. If either of these approaches correct the problem, it confirms that the problem is in the autosampler and further troubleshooting efforts along this line are worthwhile. If you haven’t done it already, replace the needle seal on the autosampler HPV, it is the culprit in many instances. If that is not effective, consider replacing the needle and sample loop.

ii. Sometimes the sample adsorbs on the sample loop because of the characteristics of sample and injection solvent. Replacing the loop with one of different composition can help solve this problem. Injection loops are available in stainless steel and PEEK. A combination of a new loop material and a different injection solvent should eliminate sample adsorption on the loop.

iii. The internal components of the HPV will become worn over time as a normal consequence of doing business. A worn rotor can be the source of carryover, but it is also usually associated with impairing the quality of chromatography peaks. Consider repairing or replacing the HPV.

8. Addressing sample-specific Carryover

a. By now you’ve tried all the easy fixes to carryover problems, so it is worth the trouble to see if the problem is specific to your sample. Sometimes you can check by examining another peak in the sample. For example, if your method uses an internal standard, try injecting a large amount of internal standard and then injecting blanks without an internal standard. Perhaps you can inject another chemically related compound that will be eluted under the current method conditions. If all else fails, you can change the mobile phase or column and run another method. If you find that the carryover problem is common to different compounds or method conditions, it is most likely a physical problem associated with the autosampler hardware or system flow path.

Reducing Carryover Using Autosampler Rinsing

As an overall rationale for addressing carryover with autosampler rinsing, use external rinsing first as it is quick and easy. If a more aggressive approach is desired, try adding internal rinsing.

External Rinse with  Autosamplers

This cleans the outer surface of the needle by immersing it and/or actively rinsing it in a rinse port before or after the sample is drawn. There are two rinse ports (immersion and flow). In simplest terms, the immersion rinse port allows you to dip the needle into a rinse solvent (R0). Commonly, R0 is mobile phase or something chemically stronger. A multi-rinse option (equipped as standard on the SIL-40C X3) allows you to use an active flow rinse port to clean the outer surface of the needle with a different combination of rinse solvents (R1 and R2). The R1 and R2 could set up a rinsing sequence of strong solvent (R1) followed by a weaker solvent (R2). The multi-rinse option may also use an active rinse solvent (R3), which controls cleaning with an active diaphragm pump.

Internal Rinse with  Autosamplers

This function can rinse the inner surface of the needle and sample loop, and the injection port. Use of the multi-rinse kit enables rinse with up to 3 types of solution (R0, R1, and R2). The strong/weak rationale should be used when setting choosing the chemistry of the rinse phase solvents. The following table summarizes available rinse options.