PhyTip Column Resins

How do I know if I am using the right antibody affinity resin? For example, I am using a mammalian expression system and isolating mouse IgG3? What resin should I use?

PhyNexus is support is available to help choose the correct resin for a particular application. When in doubt, call us! In the example posed in the question, ProA may not be the optimal resin for isolating mouse IgG3. We recommend using ProPlus or ProPlus LX resin which has a stronger affinity for this particular protein over ProA resin. In addition, Mammalian expression systems frequently do not produce very large quantities of the desired protein, so choosing a resin with a higher affinity may allow you to capture more of the product.

How long can refrigerated Affinity PhyTip columns be stored?

Each PhyTip column is manufactured and shipped with a special bead coating liquid to maintain bead hydration and prevent contamination. The tips are guaranteed to remain stable for at least six months after delivery. They will remain stable, hydrated and usable for many more months.

If I let the Protein A PhyTip column dry out while I’m using it, will the captured protein be affected?

For optimal recovery and to avoid protein denaturing, PhyTip columns should remain hydrated through the entire purification process. But there is little danger. The columns are shipped with a coating liquid to prevent the resin bed from drying. Depending on the resin, and in some cases, column styles, the bed may be translucent, partially opaque or completely opaque. The columns may be stored at room temperature in robotic systems for hours or even days (even weeks) with no danger of column damage. Once the purification process has been initiated, the columns will not dry out if used under recommended operating conditions.

What is the IMAC resin made of and why does my IMAC resin have a blue color?

A PhyTip column with IMAC resin is made of Ni-NTA (chelating) functional group bonded to an agarose resin substrate. The nickel cation, held by the NTA (nitrilotriacetate) chelating group, has an affinity for 6-HIS tag portion of a recombinant protein. Generally, there are two basic IMAC resins available: a nickel based and a cobalt based system. Cobalt resins have a slight pink/red tinge to the resin. The Nickel resin with a bluish color tinge and is the most common type of IMAC resin.

Getting Started: Sample and Column Preparation and Use

Why are PhyTip columns shipped with some fluid in them? What is the fluid?

PhyTip columns are delivered with glycerol shipping fluid in order to keep the resin bed hydrated which is necessary for optimal performance. The PhyTip columns may be used straight out of the box without preconditioning. However, the first step of a programmed method is usually an equilibration step to ensure the capture step that follows with under identical conditions column to column. The equilibration step is usually performed with a buffer that is similar to the sample being processed.

Is it necessary to condition the PhyTip columns before the capture step?

It is not mandatory but recommended in most cases. The equilibration step dilutes the glycerol that is used to pack the resin. The pH of the resin/solution is optimal for the subsequent steps.

How clear or clean should my sample be before I begin to use a PhyTip column?

PhyTip columns have been used with a variety of different sample media. In all cases, we recommend that the solution to be purified is clear of any particulate substances. In order to guarantee this, we recommend the sample is centrifuged and then the clear supernatant processed by the tip. Care should be taken not to disturb the plug of spin down residue while sample is processed. After centrifugation, it is recommended to transfer the sample to a fresh well before processing.

What is the maximum sample volume I can process with my PhyTip column?

For the 1000+ tips we recommend a maximum of 1 mL sample and for the 200+ tips a maximum of 200 μL sample. Larger samples can be processed by aliquoting sample into multiple wells of a plate which are then processed sequentially. For example, a researcher purifying 5 mL of sample is recommended to perform 5 separate capture steps. First, aliquot the protein sample into five separate 1 mL volumes. Then program the robot to perform a capture step on each of these individual aliquots. In this way, there is a maximal capture of all of the protein in the original 5 mL. The subsequent wash and elution steps can proceed as normal. This general process may be adapted to different sample volumes as desired.

How do I know if my resin is capturing protein? How do I know if my resin is saturated with protein?

If used correctly, PhyTip columns can be loaded with a surprising amount of protein.
Nevertheless, in some cases the resin may have the incorrect selectivity and no protein may be captured. In other cases, protein is captured well loading up the resin, but not all of the protein is collected from the sample volume. This is difficult to observe directly as most proteins are colorless and only after the fact can capture information be determined. Because of parallel operation of PhyTip column, the user is in complete control. Protein control standards may be run in parallel with the sample to ensure correct operation and purification chemistry. In a broader experiment researchers will retain some of the original material, the “flow-through” from each step (this is the material left after a step) so that a gel can be run at the end of the experiment to observe not only the end product – the purified, concentrated protein, but also if any of the protein of interest came through in the flow through. Reviewing this information allows the researcher to determine the success of the capture, wash and ultimately the elution steps.

How do I program my robotic system? For example, can I elute with an elution volume that is less than 3 x of the resin column bed volume?

Various default method protocols are provided by PhyNexus and excellent results can be obtained with no method modifications. However, it is possible to make programming adjustments or even major adjustments to make a new method. For example, it may be desired to increase the concentration of the recovered protein by with only 2 x the resin volume rather than the normal 3 x volume. Or larger or multiple elutions may be desired to ensure complete recovery of the protein. If programming is method development is not available within company resources, see your PhyNexus representative for advice and help with robotic programming.

Why is flow rate important when operating the PhyTip columns?

A critical component of the capture step requires that the target sample protein have sufficient time to bind to the affinity resin. By using a recommended slower flow rate, maximum capture of the desired protein can occur because this increases residence time of the sample protein with the resin matrix. The default protocol utilizes a residence time of 6 and 18 minutes for 200
μL and 1000 μL samples, respectively. The capture efficiency can also be increased by increasing the number of capture cycles.

Optimization and Enhancement of the Purification Process

What is affinity purification?

Affinity resins are designed to be selective to a particular protein tag or protein property or characteristic. Recombinant proteins are normally purified using a tag on the protein and an affinity resin that is selective to the particular tag. HIS tagged proteins are purified on IMAC affinity resins. Antibodies are purified with Protein A or Protein G affinity resins. There are many other types of affinity resins including other tag selective resins or ion exchange resins.
Proteins in a pH environment have a specific charge and can be purified with ion exchange affinity resins. The combination of affinity and specificity has been exploited to generate straightforward affinity purification methods.

What are the basic parameters that can be optimized and enhanced in the PhyTip column process?

Depending on the goals of the researcher, parameters that can be optimized or enhanced include concentration, mass, purity, or activity of the recovered protein, the speed of the purification process and the ability to accommodate various starting sample volumes. The reader is invited to read the Tip Concentrating Effect section for additional details on improving recovered protein concentrations. Some addition information is included in the questions and answers below. Your PhyNexus representative is always for further information. We are happy to help.

Why and how should the capture step be enhanced?

In order to obtain optimum recovery, whether it is concentration or mass amount, it is first necessary to make certain the capture is optimized and as much protein is captured onto the resin bed as possible. Of course this starts with the expression of the protein – it can’t be captured if it isn’t there in the first place. The best way to measure expression of a protein is to run a control protein in parallel with the sample under exactly the same conditions.
Parameters that increase capture include increasing the number of capture cycles and slowing down the capture flow rate. In many cases, adding salt to the sample solution will increase uptake.

Salt reduces the solubility of the protein and can increase the selectivity of the resin for the protein. These salts enhancing protein selectivity for the resin follow the Hofmeister series: increased selectivity effect of anion to decreased selectivity: PO43- > SO42- > COO- > Cl- ; increased selectivity effect of cation to decreased selectivity: NH4+ > K+ > Na+. But many researchers simply use NaCl in the capture with good results. Also, it is important to check the sample pH – uptake of protein is optimized at neutral pH for most affinity resins.

How does washing the resin affect recovery and purity?

When the resin is loaded, both the desired protein drug and impurities are adsorbed on to the column. As the separation method proceeds the buffer strength is increased to wash the resin. Impurities ideally completely desorb from the resin while the material of interest is still completely bound to the resin. Depending on how tightly bound to the resin, the stringency of the wash may be increased to remove as many impurities as possible without removing the desired material and without harming or denaturing the desired material. Pure desired material remains on the column. Finally, at the elution step, the buffer strength is increased so that the desired material completely desorbs from the resin and can be recovered.

In practice, wash conditions, are carefully selected so that desired product remains on the column while undesired materials are removed and washed away. If the target bio molecule to be recovered is a relatively small molecule, the binding is usually relatively high. Tagged recombinant proteins or protein fragments generally have a single point of attachment to the affinity resin. An example is HIS-tagged proteins or fragments which have affinity for IMAC resins. Smaller molecules that have a single point of attachment bind tighter to resin than a larger molecule due to steric hindrance or other factors. On the other hand, if the number of binding sites increases with size, then the selectivity may increase with size, again depending on steric hindrance and other factors. Stringent conditions can be chosen for washing without fear of removing the target bio molecule. With stringent washing, non-specific materials are efficiently removed prior to elution of the pure target material. However, strong elution conditions may be needed for complete recovery.

If the sample bio molecule is relatively large, it may not bind as tightly to the stationary phase. The stringency of the wash is normally reduced to prevent loss of the target material. Conditions for washing are carefully chosen to remove as much non-specific material as possible while still retaining the target material. However, elution is relatively easy and high recoveries from this step are possible, provided sample material was not lost in the wash step. Thus, conditions for capture, wash and elution are developed for each case to optimize the mass recovery, concentration and purity of the bio molecule.

Why and how should the elution step be enhanced?

Removal (elution) of the protein from the resin in the PhyTip column can be surprisingly difficult for some sample proteins. Running a control protein in parallel with the sample under exactly the same elution conditions is an excellent measure of elution efficiency, provided the control protein is similar in structure to the sample protein. Large bulky proteins tend to elute easily from the column whereas smaller proteins are more difficult.

Standard elution volumes are 3x of the resin bed volume. A higher concentration of protein can be achieved by reducing the elution volume to 2 x of the resin bed volume. Many researchers will perform a second elution and combine the fractions. This will increase the mass recovery but will reduce the overall concentration of the recovered protein.

How can the highest concentration of recovered protein be achieved with all protein in the sample recovered?

First, use a column with sufficient capacity to capture the entire amount of protein. Passing the sample over the resin bed several times allows as much capture of the protein from the sample as possible. Using a slower capture flow rate also increases the residence time of the sample in the resin bed which increases the capture of the sample. In cases where the sample volume is significantly larger than the total volume of the PhyTip tip, it is better to use more capture cycles rather than a slower flow rate so that eventually all of the sample protein passes though the resin bed and is captured. Better still the sample volume may be split into several volumes
and each sample volume captured in succession.

In some cases the type of resin can be important. For example, Pro A resin has a higher capacity than Pro G resin for many types of antibodies. Also, use sufficient sample target protein so that all of the resins functional groups have captured protein attached. If necessary, follow all of the optimization procedures for number of capture cycles and flow rate. Finally, using a smaller elution volume increases the concentration of the eluted protein.

How can a normalized concentration of recovered proteins be achieved?

Recovering a normalized concentration of protein (the concentration of protein recovered from the tip is uniform regardless of the starting sample concentration) can be useful in many cases.
Many assays, especially cell based assays, require a minimum and uniform amount of purified protein. This can be difficult to achieve if the expression efficiency or effectiveness of the protein is unknown or known to vary for a series of proteins. PhyTip tips are unique in their ability to normalize concentration of purified protein to a predictable level. Starting sample protein concentrations that vary widely can still be brought to a uniform concentration in the purified form. The resin bed is overloaded even with the lowest expressed protein by using sufficient sample concentration and volume. The first step is to use the smallest possible bed volume that is appropriate for the assay. In most cases, this will be the 5 μL bed volume. Overload the resin bed by a factor of two or more. If the concentration of a sample or series of samples is unknown, then it is best to process enough sample volume so that the lowest expected expressed sample has sufficient volume to overload the column and then treat all samples in the same way. It may be necessary to process several aliquots of the sample in series to overload the column.

How are bio molecules taken up and eluted from affinity columns and other columns?

The interaction of bio molecules with most chromatographic phases can be characterized by sharp isotherm “on-off” chemistry. Under any set of buffer conditions the bio molecule is either adsorbed to the chromatography material or desorbed from the material. This appears to be true for ion exchange, ion-pairing, affinity, hydrophobic interaction, reverse phase, hydrophilic interaction and normal phase material. However, this controlled interaction can only be used when the interaction of the bio molecule and column come to equilibrium under the specific buffer conditions.

What parameters are important for purified protein suitable for structural analysis?

Several parameters are important. High concentration for structural analysis is important of course, but the recovered protein must also be in the suitable buffer environment for forming crystals and have the correct folding and protein activity. Therefore it is necessary to follow the guidance above, but also to perform the operations in parallel to screen different buffers.

A two-step process for achieving ultra-high protein concentration is possible with the PhyTip tip but is beyond the present discussion. Contact Phynexus customer support for more details.

How can the highest mass amount of recovered protein be achieved?

Use the largest resin bed volume so that the resin bed is not overloaded (there is no protein in the capture flow through). The capacity of the resin bed is proportional to the bed volume. If the sample volume is large, be sure to process (with sufficient cycling) an aliquot of sample completely with each capture cycle and then repeating the cycling process with another aliquot until the entire sample is processed. Another strategy is to process only part of the total volume with each capture cycle but use several capture cycles. Eventually the sample protein will travel through the bed and be captured. Typically 10 or more capture cycles is used in this strategy. If the resin bed capacity is severely overloaded, two PhyTip columns may be used. Generally two elution aliquots should be used to recover the entire amount of protein from the resin bed.

How can the highest purity of recovered protein be achieved?

Use progressively higher concentrations of the wash solvent to remove nonspecific bound protein and check the purity of recovered protein using gel electrophoresis. The wash can remove the captured protein as well as nonspecific bound proteins and other materials. This is especially important in PhyTip column extractions because the ratio of wash solvent volume to bed volume is extremely large. So this process should be done carefully using only lowest concentration of solvent necessary for cleaning. At least two aliquots of wash solvent are necessary. More aliquots may be used but usually are not necessary.

How can the highest activity of recovery protein be achieved?

It is important that activity of a protein is reduced by denaturing or other means. A unique aspect of PhyTip columns the absence of high surface area frits that can harm proteins. But it is also important to use buffer conditions that are not denaturing in any of the processes including washing and elution. Generally this means that the buffer pH is kept neutral and at high ionic strength. Many researchers purify in parallel and screen expression conditions and purification conditions. Activity can be tested with enzymatic assays, ELISA, SPR or a functional test.

How can the purification process be shortened?

One answer is to process samples in parallel as much as possible. Of course reducing the number of cycles will reduce time. Often just 2 or 3 cycles are sufficient for the capture process. But use fewer capture cycles or use a fast flow rate but only if necessary. Usually a larger bed volume column will capture the limited sample in a shorter time period. If possible reduce the wash and elution/enrich cycles before the number of capture cycles is reduced. Often, for washing, usually only 1 cycle of each solvent is adequate. Lowering the number of capture cycles down to 2 cycles is acceptable in almost all cases but in some cases this could lower the amount of protein captured and increase the deviation of the recovery from sample to sample.

How can the smallest sample volumes be processed?

The ability to process small sample volumes enables the possibility of the expression of small volume of proteins, reducing the amount of material required to produce the desired protein.
Normal sample volumes range from 0.2 mL to 1 mL, but samples can be as large as several milliliters or as small as a few microliters. Because of the low dead volume and low resin bed volume of the PhyTip tips, very small elution volumes can be used to elute protein from the resin bed. Very high protein concentrations are achieved with these extremely small elution volumes. The concentration of the protein increases by using decreasing elution volumes (although less mass amount of the protein may be recovered). Use the smallest resin bed volume possible and use the smallest elution volumes (the concentration increases with decreasing elution volume).

How should large sample volumes be processed?

Two different methods are normally employed. Sample volumes larger than the tip chamber volume can be processed simply by inserting the column in the sample and processing with sufficient number of cycles so that any sample protein in the sample has a chance to contact the resin bed. A general rule is to double the number of capture cycles exponentially for every chamber volume of sample. Thus if a 1 mL sample in the 1000+ column (with a 1 mL sample chamber) uses 4 capture cycles, then a 2 mL sample using the same column should use 8 capture cycles, a 3 mL sample should use 16 capture cycles and so on.

A better method is to split the sample into as many aliquots as necessary and process each aliquot in succession until the entire sample is captured. Then proceed on as normal with the wash/purification step and the elution/enrichment step.

What factors are important in PhyTip sample capture?

Of course the affinity chemistry must be suitable to selectively capture the protein of interest.
For example, Protein A resin will capture Mouse IgG2a but will not capture Mouse IgG1. Refer to the Protein A and Protein G affinity information and product insert sheets to identify the correct affinity resin for a particular type of antibody capture. ProPlus and ProPlus resins have selectivity that combine Protein A and Protein G. IMAC affinity resin is used to capture HIS tagged recombinant proteins. The sample pH must be correct – in most cases, a sample of pH 7 is desirable for capture. For some sample proteins, it is important to increase the column bed residence time to compensate for low-affinity interactions. Capture residence time is increased by decreasing capture flow rate and/or increasing the number of capture cycles.

What is the maximum loading capacity for a PhyTip Tip? For example, what is the loading of a Protein A PhyTip column?

When using the largest bed volume 1000+ PhyTip tips, up to 2 mg of human IgG can bind to the
Protein A affinity resin provided the sample contains enough protein and proper loading procedures have been followed. To fully load the resin bed, at least a slight excess protein is needed in the sample. High loading is achieved with slower flow rates or more loading cycles; however, if the sample protein excess is large, flow rate and number of loading cycles are less important. Contact your PhyNexus representative for further information.

What size proteins can I purify with the PhyTip system? For example, can I purify protein complexes that are very large?

Most proteins that researchers study are in the 5 to 200 kDa size range. Interestingly, as the protein size increases, the larger, more bulky proteins may not stick as tightly to the affinity resin and the conditions for purification are usually less stringent to prevent loss of protein.
Nevertheless, these proteins are purified quite easily. Protein complexes however are much larger e.g. 1 MDa to perhaps the 8 MDa range. Proteins in a complex held in a strong core of proteins (proteins are tightly associated with each other) are usually captured by packed bed columns and will also be captured by the PhyTip columns. Transient (non-core) proteins are often difficult to isolate in packed bed systems; they are often too large and fragile to survive the process of flowing through the packed bed. The gentle action of the PhyTip system will enable purification of transient proteins in protein complexes. Each system should be examined on a case by case basis.

What are affinity tags?

Recombinant proteins are generated by introducing an extrachromosomal DNA vector (plasmids containing the gene of interest) into cells. Then the plasmid vector utilizes the cell’s machinery to express the gene to produce the protein of interest. Affinity tags attached to the
N-or C-terminus of the protein are expressed along with the recombinant protein. These affinity tags make it possible to perform affinity purification with the appropriate affinity resin. After capture and column washing, the recombinant protein is recovered by eluting the column with a buffer solution. In some cases, tags can be enzymatically cleaved of the column. An example of this is engineering TEV sites into the DNA vector in addition to DNA expressing the tag. The affinity resin will capture the protein with the tag while leaving the TEV site available for cleaving. Some tags can be attached to proteins directly (without plasmid vectors). An example of this is the chemical attachment of the biotin tag to lysine residues on the sample protein.

What are some common affinity tags and their counter ligands?

6-Histidine peptide residue (6-His) tag will bind to IMAC affinity resin containing divalent metals including Ni(II) or Co(II). Glutathione S transferase (GST) tag will bind to glutathione (GST) affinity resin. Biotin tag will bind to Streptavidin affinity resin. FLAG tag will bind to anti-FLAG antibody affinity resin. Maltose Binding Protein (MBP) tag will bind to amylose affinity resin.
Myc tag binds to Anti-Myc antibody affinity resin. Calmodulin Binding Protein (CBP) tag binds to Anti-calmodulin binding protein antibody affinity resin. HA peptide tag binds to Anti-HA antibody affinity resin.

What are the major challenges to affinity purification?

Affinity purification methods often struggle to maintain or increase the concentration of the protein being purified while at the same time providing a protein that is pure and active. This is especially true as the amount or volume of protein being purified is decreased. It can be very difficult manipulating microliter volumes of sample. Finally, it is difficult to routinely purify large numbers of samples in parallel in a laboratory environment. PhyTip column technology is designed to easily overcome all these challenges.

Protein Recovery, Purity and Activity

If my antibody is very dilute (e.g. 5 μg/mL) and I have 5 mL of solution, which PhyTip columns do you recommend and how do I recover as much protein as possible?

Using the 1000+ PhyTip columns with the 20 μL bed, divide the 5 mL sample into 5 equal volumes of 1 mL. Perform the capture step for at least 2 cycles, preferably 4 cycles, sequentially for each 1 mL of sample. After column washing, elute the protein captured with 3x the resin bed volume of elution buffer. In an alternate capture procedure, the sample could be put into at least a 6 mL vial and the entire sample processed. In this case, only a portion of the sample can travel through the column in any one cycle. Approximately 15 cycles should be used to ensure that the sample protein travels through the column at least once and is captured. After sample capture, process the PhyTip column in the normal manner as described above. The first sequential procedure is preferred.

I followed all of the instructions for isolating my protein, but I see nothing on my SDSPAGE gel, what do I do now?

Check for the presence of your protein in the capture flow-through solution using SDS-PAGE. If none is present, then there are only two possibilities. Either the sample has not expressed and there is none to be captured or the sample did express and was captured but did not elute from the column. Highly expressed samples can sometimes be detected directly on the SDS-PAGE.
Assuming that sample has expressed, then confirm that the sample capture pH is correct
(approximately 7.4) and that the sample is compatible with the column chemistry being employed. Also, increase the elution strength with parallel experiments to study the effect on recovery. Many users use a known standard protein with known characteristics. The sample and standard are processed in parallel whenever a new unknown protein is being expressed and/or purified for the first time.

I am using the PhyTip ProA tips to isolate Mouse IgG, but I don’t see any bands in my gel.
How do I know that my system is working?

Review the expression system and the type of mouse IgG in the sample. Refer to Protein A and
Protein G affinity information sheets for reference tables of antibody selectivities for affinity resins. Check the flow-through from your capture steps to verify that the protein was captured.
Check the wash conditions to make sure the specific protein of interest was not removed in the wash step. Next, check to see if the protein is still attached to the resin after the final elution step (pH of the buffer may not have been low enough). Finally, check to see if the eluted protein neutralized effectively and did not denature. Many users run an antibody standard in parallel with the sample to help monitor loading and recovery performance of the sample.

What if I put excess protein through the PhyTip column. Does that affect my ability to capture the protein I want?

No, once all capture sites are full, excess protein does not affect the existing captured proteins.
This process will produce an extremely concentrated purified protein when the sample is finally recovered in the small elution volume.

What is the maximum mass that I can get from a 20 μL ProA, ProG and IMAC column?
What about other bed volume columns?

The PhyTip process can often capture more protein onto the resin bed than suggested by the original resin manufacturer. The repeat cycling of the sample capture process used by the
PhyTip column maximizes resin performance. Multiply the resin capacity per unit volume by the number of μL in the column bed.

Can I recover more material if I do two elutions rather than one?

Experiments suggest that with a single elution step (and depending upon the affinity of the resin and the elution strength of the elution buffer) anywhere between 60 and 80% of the purified protein can be removed in a single elution step. This percentage can be increased with a second or third elution step if the end result is to recover more mass. But this will be at the expense of concentration if the elution volumes are combined.

How do I know that all of the protein has been eluted from the resin?

Elution of protein from the tip can produce two different end results. If the requirement is for highest concentration then the minimum volume of elution is recommended. However, this may result in a lower percentage of purified protein being removed from the column. If the goal is maximum mass of purified protein, then several wash steps will ensure maximal removal of protein from the resin. This however will reduce the concentration of final protein. It is unlikely that 100% of protein will be removed from the resin, and researchers can expect that with sufficient elution, approximately 95% of protein can be removed from the resin.

I have heard that you can elute from an IMAC resin using variable pH. Is this true?

Yes, low pH elution buffer can remove a His-tagged protein from an IMAC resin. In addition, adding a chelator such as disodium EDTA to the elution will remove the protein from an IMAC column but will remove the nickel metal as well.

My IgG protein doesn’t seem to be pure. I expected to see only one band but I get two on the gel. What’s wrong?

One would see two bands on SDS-PAGE: a light chain and heavy chain. A single band is expected if the sample has not been treated to break the sample into the 2 chains.

I am running an IMAC resin and my final product is not as pure as I had hoped. How can this be improved?

Wash buffers for IMAC resin will often contain low levels of Imidazole that help wash away some of the nonspecific 2, 3, 4 or 5 His tagged variants that may have been expressed along with the specified 6-His product. Imidazole in the wash helps remove any histidine rich protein in your sample that competes for Ni(II) in the resin. Sometimes the concentration of the imidazole is not high enough to remove these contaminants (the wash is not stringent enough) so the final product will not appear as pure as it could be. But care must be taken if the wash buffer strength or stringency is increased because the sample protein of interest might also be removed in the wash. If the wash stringency is too high, the protein might be very pure, but the yield might be very low.

Why do you recommend two different types of wash buffers while using ProA or ProG resin. What is the second buffer and what does it do?

Two wash buffers enhance the elution step by reducing the pH shock when moving from a pH 7 wash buffer to a pH 2 elution buffer. Low pH elution buffer has higher strength to break the binding between resin and protein and can improve protein recovery. If there is no intermediate saline wash, the residual pH 7 wash buffer that resides in the resin will decrease the pH strength of the elution buffer which then results in a lower yield of eluted protein. Adding an extra step to the process removes the buffering effect of the first wash step and enables the low pH of the final elution step to be effective and produce the highest recovery of purified protein. The second wash can be very important to achieving reproducible elution of the protein and is always recommended.

What is the minimum recommended wash buffer volume that still achieves the highest purity of sample?

This will depend upon the volume of the resin in the tip, the type of protein being purified, and the types of contaminants that are mixed in with the protein of interest. For example, a 200 μL wash of a 5 μL resin bed will probably produce the optimal purity results. It is possible that the same level of purity could be achieved using a 100 or even a 50 μL wash volume, but given the cost of the wash buffer and the difference in time between a 50 and a 200 μL wash, the recommendation is to go for the highest volume rather than try to compromise.

Is imidazole in the equilibration buffer and sample buffer? If so, why?

We recommend using the following equilibration buffer while using IMAC resin: 50 mM sodium phosphate, 1.5 M sodium chloride, 25 mM imidazole, pH 7.4 Note: This is a 5X buffer and needs to be diluted to 1X with water before use. Imidazole is present to clean the column from any material that might bind to Ni present on the column. Imidazole in the sample buffer can help keep the protein pure by limiting the capture of impurities along with the recombinant sample protein.

What is the difference between high and low stringency buffers?

Low stringency washes remove weakly bound nonspecific impurities. If the sample protein is large and bulky, then the column should be washed with low stringency buffers to avoid losing sample protein. Higher stringency washes can be used for tightly held sample proteins. For IMAC resins, the difference between the low and high stringency buffers is the concentration of imidazole used. For IMAC, a low stringency buffer is 5 mM imidazole and a high stringency is 20 mM imidazole. For ProA/ProG resins, the stringency of the wash can be controlled by the volume of the wash. Smaller wash volumes provide a low stringency wash. Large wash volumes provide a higher stringency wash. Stringency for these resins can also be controlled by the wash pH (lower pH is a higher stringency wash), but this should be done carefully with testing to avoid sample protein loss.

What is the relationship of high protein recovery and high protein activity?

Interestingly, high protein recovery and high protein activity are not necessarily correlated showing the need for method development. Using the PhyTip columns operated in parallel, it is quite easy to study two or more variables, such as the effect of pH and salt concentration, in the same set of experiments. The complex interactions of sample and contaminants with the resin can be studied to optimum yield.

In protein purification, not only is yield important but the recovered protein must remain active and not agglomerated. In one experiment of parallel method development for purification of an antibody on one type of Protein A column, different conditions of capture, wash and elute (8 each) were tested for each of 4 clones. The recovered proteins were screened for yield (ELISA) and retention of activity (SPR). The results show that high recovery and high (retained) activity are not correlated and both goals should be considered when developing separation conditions for proteins. It is difficult to consider these goals without parallel purification.

When separation conditions are established, parallel operation of columns can be used to separate up to 96 samples at-a-time.

Why do I need the neutralization buffer?

If low pH elution is used, the protein may denature if left at low for an extended period of time.
Although there is no set time period, the protein buffer pH should be raised to neutral pH in an expedient manner. The neutralization buffer is concentrated and can be added directly to the eluted protein to raise the pH to a physiologically stable environment. When necessary, confirm the pH has been raised to 7 after an aliquot of the concentrated neutralization buffer has been added to the eluted sample.

Will my purified protein still be active or will it denature?

PhyTip tips are designed to have ultra-low non polar surface area that could come in contact with proteins. Proteins may unfold or denature when they contact a non polar surface. This is especially true for membrane proteins because these proteins have a large non polar component to their structure. Limiting the amount of non polar surface in the PhyTip column helps keep proteins from denaturing. PhyTip purified proteins are likely to maintain their activity during and after the purification process.

What is the optimal elution pH for a Protein A or Protein G tip? How quickly should I neutralize the eluted protein? How should the eluted protein be neutralized?

The optimal pH will vary depending upon the protein that’s captured and the affinity it has for the resin. Smaller proteins stick tighter to the resin. Obviously, the gentlest conditions are desired for elution. Most proteins will elute from the resin at pH 2.5 – 3.5 but it may be necessary to elute as low as pH 2.0. See customer support for the types of low pH buffers used for elution. After elution, neutralization with a buffer is recommended to bring the pH back up to neutral as soon as possible – no longer than a few minutes. The neutralization buffer may be a pH 8 or 9 depending upon the strength of the buffer used to elute the protein.

High Performance Immunoprecipitation (HPIP)

What is immunoprecipitation (IP)?

IP is an abbreviation for individual protein immunoprecipitation. Immunoprecipitation (IP) takes advantage of natural antibody antigen interactions to purify a particular protein from a sample of biological origin. The process uses a specific antibody to capture its counterpart antigen protein in sample types such as crude lysates of bacteria, plants, animal tissue or other various body fluids. This method enables protein isolation and concentration to aid in the study and assay of proteins of interest that would otherwise be difficult to detect in applications such as western blotting. IP can also help identify and interrogate protein-protein interactions.

Operationally in IP, a protein mixture or cell lysate containing the protein of interest is brought into contact or incubated with a specific antibody chosen for the purpose. Antibodies specific for given proteins are commercially available from a number of companies. The “immuno” in the term immunoprecipitation refers to the antibody selectivity and mechanism for capture of the protein. The “precipitation” portion of the term refers to the capture of the complex onto a solid support. There are different ways that IP can be performed to recover the protein of interest.

What are the different ways that a PhyTip columns can be used to purify a protein of interest in an IP experiment?

There are two basic antigen/antibody capture methods used in IP. The first method, called the indirect method, involves forming the antibody/antigen complex in solution prior to isolation.
After incubation, the complex is captured by passing it through a PhyTip Protein A or Protein G column. The antibody portion of the complex attaches to Protein A or G resin in the PhyTip tip, pulling the antigen along with the antibody. After capture, the complex is washed and then eluted off the column using a low pH buffer.

One of the advantages of forming the complex first and then capturing is the time, temperature, and chemical conditions for complex formation can be easily and finely controlled. In this way, complex formation is reliable, predictable and reproducible. For example, the complex formation can be performed in a refrigerator at 4oC for a specified time with specific reagent concentrations. However, formation of the complex relies on the diffusion of reagents to make contact. Mixing by sonication can be performed in some cases. Diffusion or mixing of this type is very slow sometimes requiring hours, or even overnight processing.

Formation of the complex is an equilibrium process therefore, the concentration of the antibody and/or antigen is a factor in how rapid and to what extent the complex can be formed. Because the reagents may be dilute in the indirect method, the driving force for complex formation can potentially be low. Both of these issues of diffusion and driving force can slow formation of the complex and reduce the amount of complex that is ultimately formed. Nevertheless, because of the tip concentrating effect, the high concentrations of the final product are eluted from the PhyTip column. Further information is available in the PhyNexus white paper “High Performance Immunoprecipitation Using the PhyNexus PhyTip System.”

The second method, called the direct method, involves loading the antibody directly onto a
Protein A or G PhyTip column by back and forth flow. The conditions can be optimized for high loading of the column while still using small amounts of antibody. After the “antibody column” is constructed, the column is placed in the sample and the antigen is captured from solution, again by using back and forth flow. There are several advantages of loading the antibody onto a Protein A or G column and then capturing the desired protein. First, a controlled, high concentration of antibody is introduced to the Protein A or G resin via active transport (back and forth flow) as opposed to diffusion. Active transport (actively flowing or bringing the reagents to the reaction site) decreases the time and increases the control of the antibody so that very little reagent is used in each experiment. In effect, the researcher is making a selective resin for the next step, the capture of the antigen. Because the antibody is captured on the resin in high concentration (relative to bulk solution) and the sample antigen is brought to the antibody via active transport (back and forth flow) the capture of the antigen is rapid, predictable and efficient. After capture, the complex is washed and then eluted in with low pH buffer as normal. This procedure is another variation of High Performance Immunoprecipitation (HPIP).

Why is IP by PhyTip called “High Performance Immunoprecipitation (HPIP)”?

High Performance IP is characterized primarily by speed and miniaturization with the following key attributes:
1) Fast; Active transport drives the equilibrium of the reaction and dramatically decreases the time required to obtain high purity targets.
2) Linear; Back and forth flow pushes the equilibrium reactions to completion. Reagent concentrations can be adjusted to shift reaction equilibrium and control speed of equilibrium.
The amount of antigen recovered is proportional to the amount of antigen in the sample.
3) Sensitive; Concentrations of the recovered protein are higher than by any other method.
4) Predictable: Again, column interactions are driven to completion. Conditions and time can be controlled. If using a PhyNexus MEA, experiments can be performed in a refrigerator giving temperature control to the process.
5) Because the method is controllable and predictable, any set of conditions developed by an operator in one laboratory can be transferred to another operator or even another laboratory.
6) Parallel operation; The process is controlled and can easily be performed under exactly the same conditions.
7) Cost effective; Small volumes of resin in the tips and active transport to load the antibody make effective use of costly antibody and reagents at no cost of performance.

You mentioned that PhyTip HPIP is cost effective by making effective use of the antibody.
How is this possible?

It may seem counterintuitive, but the best results are obtained by using the smallest column possible. The Tip Concentrating Effect described in this question and answer explains that the highest concentration of protein is recovered from the smallest column. . Because a small column can be used effectively, the amount of costly antibody needed to load the column is relatively small. We recommend the 20 μL bed column for HPIP. But if an antibody is more expensive than normal, it is probably worth trying out the 5 μL bed PhyTip column and comparing the results. In many cases, the 5 μL bed will perform very well. Another interesting aspect of using the smaller bed column is the reduction of nonspecific binding. Since the Protein A or Protein G column is small, it is possible to load up virtually every site on the column. Increasing the concentration of the antibody on the column reduces the possibility that other (nonspecific) contaminants bind to the column and are recovered with the antigen.

Can HPIP be performed without using different antibodies?

Literally thousands of different antibodies are now available from perhaps a dozen companies.
However, it still may be difficult or inconvenient to acquire an antibody that specifically targets a particular protein of interest. To simplify this, researchers will engineer antibody selective tags onto either the C- or N- terminal end of the protein of interest. The advantage is that the same tag can be used time and again on many different proteins and the researcher can use the same antibody. Examples of tags that can be engineered into proteins are the Green
Fluorescent Protein (GFP) tag, Glutathione-S-transferase (GST) tag and the FLAG-tag tag. There are several examples of an antibody that are selective for a particular tag e.g. GFP that can be purchased and bound or captured by a PhyTip column. While the use of a tag to enable pulldowns is convenient, the method may not be as biologically relevant. The tag itself may either obscure native interactions or may introduce new interactions.

What are GST tagged IP and HIS tagged HPIP?

This procedure is somewhat of a misnomer but should be mentioned here because of its simplicity and power. The technology, sometimes called a “pull-down” affinity purification technique, is similar to immunoprecipitation except that the antibody function is replaced by some other affinity system. In this case, the affinity system is either a GST-tagged protein that can be captured by glutathione agarose beads or a His-tagged protein that can be captured by Ni-IMAC beads. The recombinant tagged protein acts as the “bait” to capture a specific binding partner sometimes called the “prey.” The procedures and methods of use are the same as conventional IP. For example, the His-tagged bait protein is incubated with a cell lysate and the protein complex pair is captured on PhyTip IMAC column. After washing, the protein complex pair is eluted for analysis by gel, western blot, mass spectrometry, etc. Alternatively, the His-tagged protein can be loaded on a PhyTip column and nonspecific bound material washed away. Then the PhyTip column can be used to capture the protein of interest and recover the protein.

What is a false positive?

A false positive is a recovered protein that is not actually associated with the antibody baited protein. False positives can occur through exposure of the proteins in the sample to a high surface area, non-polar surface such as a column frit, column body or resin matrix. This binding is sometimes called nonspecific binding because although the false positive protein was recovered it was not due to the protein-protein interaction of interest. However, false positives can also be due to secondary interaction of the antigen or low selectivity interactions of the antibody. Secondary interactions are related to Co-IP complex formation. Low selectivity interactions are when an antibody can bind to several different proteins (albeit the protein of interest has the highest selectivity interaction).

There is one interesting note about false positives. They can be dealt with provided the results are consistent and predictable. In other words, if a material is recovered that is found to be nonspecific to the complex but can be recovered consistently and predictably, then the nonspecific material can be reliably dismissed as being not important. In many cases, false positives can be dismissed by identification and subsequent determination of their actual function. The protein may be identified by excising the band from the gel and analyzing by mass spectrometry.

What is a false negative?

A false negative is simply when the antigen is not recovered even though it is present in the sample. This can occur because the conditions for complex formation are not correct. However, this is more likely to happen because the antigen is low concentration and the recovered protein is therefore also very low concentration. Nevertheless, the tip concentrating effect of the PhyTip column can increase the concentration of the recovered protein by up to a factor of 10 over competing technologies.

How is immunoprecipitation (IP) related to co-immunoprecipitation (co-IP)?

Immunoprecipitation is closely related to co-immunoprecipitation. In co-IP, intact protein complexes (i.e. antigen along with any proteins or ligands that are bound to it) are captured and purified by selecting an antibody that targets a known protein that is believed to be a member of a larger complex of proteins. Targeting this known member makes it possible to pull the entire protein complex out of solution and subsequently, by using mass spectrometry, identify unknown members of the complex.

Co-IP works best when the proteins involved in the complex bind to each other tightly, making it possible to pull multiple members of the complex out of solution. This process of pulling protein complexes out of solution is sometimes referred to as a “pull-down.” Identifying the members of protein complexes may require several rounds of precipitation with different antibodies for a number of reasons. A particular antibody often selects for a subpopulation of its target protein that has the epitope exposed, thus failing to identify any proteins in complexes that hide the epitope. In many cases, only less than half of the proteins of a given complex are captured with a single antibody. However, the first round of co-IP will often result in the identification of many new proteins that are putative members of the complex being studied. The researcher will then obtain antibodies that specifically target one of the newly identified proteins and perform a new co-IP experiment. Likewise, this second round of precipitation may result in the recovery of additional new members of a complex that were not identified in the previous experiment, and so on. As successive rounds of targeting and co-IP are performed, the number of identified proteins will continue to increase. The identified proteins may not ever exist in a single complex under a given set of conditions. Rather, they may represent a network of proteins interacting with one another at different times under different conditions for different reasons.

How is IP related to chromatin immunoprecipitation (ChIP)?

Chromatin immunoprecipitation (ChIP) is a method used to determine the location of DNA binding sites on the genome for a particular protein of interest. This technique provides information regarding the protein–DNA interactions that occur inside the nuclei of living cells.
The in cellulo nature of this method is in contrast to other approaches traditionally employed to answer the same questions. By using an antibody that is specific to a putative DNA binding protein, one can immunoprecipitate the protein–DNA complex from cellular lysates. Crosslinking is often accomplished by applying formaldehyde to the cells (or tissue) or by using DTBP, for example. Following crosslinking, the cells are lysed and the DNA is sheared by sonication. Then an antibody is used in the sample in the same manner as in IP to capture a protein and its associated (crosslinked) DNA fragment. After reversing crosslinks, the DNA can be identified by sequencing. The location on DNA that the specific protein binds to can then be determined by
PCR, qPCR, sequencing, or microarray analysis

Drug Manufacturing Chromatography Method Development and Continuous Chromatography

What is Continuous Chromatography?

Continuous Chromatography is a new process for manufacturing biomolecules, primarily antibody drugs. The biopharma industry is facing increasing demand for large quantities of recombinant proteins due to the growth in the protein drug market as well as advances in proteomics creating the need for faster and more rugged manufacturing processes. Historically, large scale manufacturing for pharmaceutical has relied on a single column chromatography recovery processes, but is now moving increasingly to Continuous Chromatography methods.

Continuous Chromatography is a semi-continuous process that drives a primary column to complete loading equilibrium while capturing overflow on a secondary column next in line. Subsequently, when loading is complete, the primary column is removed. The former secondary column becomes the new primary column with a new secondary column installed in series to catch overflow. The removed primary column is washed to remove impurities and eluted to recover the drug protein product. After that, the column is cleaned and conditioned to make the column ready for reuse. Typically a column resin packing is reused about 200 times and then the column is repacked with new resin.

In Continuous Chromatography, the loading of the column is based on column selectivity for the drug protein under the loading buffer conditions that are applied. Loading is not based the kinetic rate of interaction, flow rate linear velocity, packing uniformity or packing diameter. The column is loaded to equilibrium. The column is loaded to equilibrium, regardless of flow rate and break through profile.

What does “loaded the column to equilibrium” mean and why is that important?

Even if the breakthrough flow profile is not sharp, the overflow material is not lost. Overflow material is captured on a second column that is held in series with the first column. This means that the column is completely loaded because the chemistry of interaction is brought to completion and equilibrium. The column equilibrium is the result of resin selectivity and the buffer conditions that are used.

Other reasons on what this means and why it is important will be explained later in this Q&A in the discussion on Dual Flow Chromatography.

How is this different than the single column process currently in use?

In a single column process, the manufacturing column is loaded as rapidly as possible using a rapid flow rate while still maintaining a sharp sample breakthrough profile. Loading is complete when the drug just starts to break through the column. If the column breakthrough profile is sharp and the column is loaded to the resin at the outlet end of the column, this column is also said to be completely loaded and at loading equilibrium. After loading, the column is washed, eluted to recover material and then reconditioned to start the process again.

For drugs that have a rapid kinetic uptake (and a sharp, flat breakthrough curve) high flow rates may be used. But drugs with slow loading kinetics, the capture profile may not be sharp and product could be lost in the overflow. In these cases, flow rate must be lowered to sharpen the breakthrough curve, albeit at a penalty of increased manufacturing process time.

So loading to completion and to equilibrium is possible with a single column system. However, in these cases, the flow control is important to maintain a sharp profile. The velocity required will vary with the kinetics of capture of the individual proteins of interest.

Why is the market shifting?

Since the capture kinetics of every protein is different, optimizing a method to manufacture a drug with a single column takes rigorous development of the flow rate and close monitoring of the column during operation. Failure to do this results in valuable product being lost.

It is now possible to err on the side of having too fast flow rates without losing product. In addition, capture is now a continuous process – there’s no interruption. Both of these vastly increase the speed of manufacturing. Productivity has essentially more than doubled.

But isn’t it more expensive to use a continuous process?

Capital hardware costs are higher. Yes, more switching and control hardware and software are needed. More manufacturing columns are needed. But column hardware costs of the individual columns can be decreased because smaller columns can be used.

The overall operating costs, including the resins costs, are identical – it is not more expensive. No additional resins and buffers are needed, so there are no additional consumable costs per unit of delivered product.

The labor costs are lower because productivity of producing the final product is higher.

Are there any other benefits to Continuous Chromatography?

Yes, it turns out there are several somewhat unexpected benefits. At the conference, PhyNexus introduced a new chromatographic concept: Dual Flow Chromatography. The paper presented was “Dual Flow Chromatography for Parallel, Automated, High Throughput Process Development”

Dual Flow Chromatograph (DFC) is a process where separations are performed with back-and-forth flow of the samples and the mobile phase through pipette tip based columns. Typical column bed sizes are very small, ranging from 5 to 160 µL. Although different than conventional unidirectional flow-through chromatography, chromatographic principles still control the dual flow process. Capture of sample is brought to equilibrium with back and forth flow through the column; the flow is the sample as pipetting the solution several times. The back and forth cycle is completed until capture is complete – usually in 4 to 6 cycles. Separations are performed 96 at-a-time using standard laboratory liquid handing robots.

How does Dual Flow Chromatography benefit Continuous Chromatography?

Although the flow dynamics are different, both Continuous Chromatography and Dual Flow Chromatography operate on the principle of bringing column resin and sample molecule interaction to equilibrium and completion. In both methods, sample molecule capture is complete based on resin and buffer conditions, regardless of sample molecule kinetics.

But how does that make it beneficial?

First, because it makes it possible for Dual Flow Chromatography to be used to predict the quantity and the quality of a product that can recovered from a column regardless of the column size. Small columns can be used to predict the performance of preparative and manufacturing columns. In the paper presented, a plot of amount of protein captured vs. the column size was linear with a slope of one. Scaling can be accomplished regardless of column size, provided the columns are completely loaded. This is will be discussed further in the Q&A.

Second, because the separation conditions developed in parallel under automated conditions can be used to predict the performance of a continuous chromatography column operated under the same buffer conditions. Multi-variable conditions can be tested quickly and compared directly. Literally thousands of methods with different resins, capture, washing and elution conditions can be tested.

Existing method development & optimization techniques such as plates and FPLC are not keeping pace with advances in manufacturing scale production. Lab scale method development is slow, and focuses primarily on protein mass capture (yield) and, to a lesser extent, protein purity.

Lab processes are not designed for efficient multi-variate, multi-parallel experimentation for yield, purity and protein activity. Lab processes may not include protein activity or agglomeration measurements – an absolutely critical measure of ultimate effectiveness at large scale.

Why is multi-variate screening important?

Proteins are complex molecules. Agglomeration of proteins can occur at any step in the purification process. Proteins may denature. Recovery must be high. The proteins must remain denatured and active. Multi-variate screening of conditions including buffer type, pH, concentration, salt additives, surfactants, etc. is the only possible way all these goals can be achieved.

Dual Flow Chromatography can be used to develop and evaluate literally thousands of possible manufacturing procedures within a matter of days – all done in miniature columns and processed in parallel on automated robotics.

In one DFC experiment described in the paper presented, 288 purifications conditions were screened for 4 clones – 72 different procedures for each clone. The work was completed within a day. Yield was measured by ELIZA and protein activity measured by SPR. The result showed that it is possible to have high yield and very low activity. But success is difficult and only if the correct conditions are used. In fact according to these experiments, it is much more likely that only high yield is achieved or high activity is achieved, but not simultaneously.

Other references where DFC multi-variant method development was used for scale-up: R. Hopkins, D. Esposito, and W. Gillette “Widening the bottleneck: increasing success in protein expression and purification” J Struct Biol. 2010 October ; 172(1): 14–20., M. D.Wenger, P. DePhillips, C. E. Price and D. G. Bracewell “An automated microscale chromatographic purification of virus-like particles as a strategy for process development” Biotechnol. Appl. Biochem. (2007) 47, 131–139 and Michael Rauscher, John Welsh, Jennifer Pollard Process Development and Engineering, Bioprocess Development, Merck & Co., Inc., Kenilworth, NJ, USA “Application of High Throughput Resin Tips for Chromatography Development” poster PREP 2015, The Loews Hotel, Philadelphia, July 26-28, 2015.