text:
Proteomics/Protein Separations - Chromatography/Ion exchange

Chapter written by: Laura Grell and Alexander Butarbutar Contact llg3875@rit.edu or nbb3924@rit.edu for contributions Chapter modified by Kai Burnett and Dalia Ghoneim Contact kab9783@rit.edu or dxg6098@rit.edu

Ion Exchange Chromatography
Ion exchange is probably the most frequently used chromatographic technique for the separation and purification of proteins, polypeptides, nucleic acids, polynucleotides and other charged biomolecules (Ref1). The reasons for the success of ion exchange are its widespread applicability, its high resolving power, its high capacity and the simplicity and controllability of the method (Ref2)

Just as with other forms of chromatography, ion exchange chromatography utilizes both a stationary and mobile phase. The stationary phase in this method carries either a positive or negative charge. The charged stationary phases are named according to the types of charged particles that bind to them.

The stationary phase for anion exchange consists of a synthetic polymer (resin) which contains many positively charged functional groups, typically quaternary ammonium ions. Negatively charged species are attracted to the resin and travel more slowly down the column. Similarly, positively charged species are attracted to a cation exchange resin's negatively charged functional groups. Sulfonate groups are commonly found as side-chains in cation-exchange resins. The stationary phases are available as granular material or beads, but prepacked column are now readily available : GE HealthCare, Sigma-Aldrich, Rohm and Haas, Sybron Chemicals, Edvotek Ion Exchange Chromatography Kit, Bio-Rad. The particles that bind to the stationary phase can be released by changing the buffer conditions.

Charge On Protein
The charge on the protein affects its behavior in ion exchange chromatography. Proteins contain many ionizable groups on the side chains of their amino acids including their amino - and carboxyl - termini. These include basic groups on the side chains of lysine, arginine and histidine and acidic groups on the side chains or glutamate, aspartate, cysteine and tyrosine. The pH of the solution, the pK of the side chain and the side chain’s environment influence the charge on each side chain. The relationship between pH, pK and charge for individual amino acids can be described by the Henderson-Hasselbalch equation, which is described in detail elsewhere:

In general terms, as the pH of a solution increases, deprotonation of the acidic and basic groups on proteins occur, so that carboxyl groups are converted to carboxylate anions (R-COOH to R-COO-) and ammonium groups are converted to amino groups (R-NH3+ to R-NH2). In proteins the isoelectric point (pI) is defined as the pH at which a protein has no net charge. When the pH > pI, a protein has a net negative charge and when the pH < pI, a protein has a net positive charge. The pI varies from protein to protein. This is the reason ion exchange is useful for separating proteins. If a buffer containing more than one protein is used with an anion exchange resin, then the most negatively-charged protein will be most attracted to the stationary phase and will therefore elute last and the protein with the highest positive charge will elute first.

Stages in Ion Exchange Chromatography
The Ion Exchange process can be separated into four basic stages (Res1):

 * 1) Equilibration
 * 2) Application of sample
 * 3) Elution
 * 4) Regeneration

Equilibration
This stage involves setting up the desired starting conditions so that the system is ready for the ion exchange process. In this step the a buffer with the desired conditions is applied, and all of the charged group in the stationary phase are associated with ions of the opposite charge. When this process in complete, equilibrium has been reached.

Application of Sample
In this step, the sample is applied to the stationary phase. Only proteins carrying a charge opposite to the stationary phase will bind to it while those with the same charge or no charge will not bind. These unbound particles will wash out during this stage. In the chromatogram above, protein A never binds to the stationary phase and washes out forming the first peak.

Elution
This step involves changing the buffer conditions to particles that have bound to the stationary phase. There are several different ways to do this. One option is to change the pH of the buffer solution. As mentioned before, when the pI of a protein is the same as the pH of a solution, the net charge of the protein will be zero. Therefore, when the buffer solution's pH reaches the pI of the protein, the protein's net charge will be zero. The protein will no longer bind to the stationary phase and will be released and washed out. Another method to elute the bound protein is to increase the salt concentration. As the salt concentration of the buffer increases, salt ions replace the bound protein. Proteins with weaker ion interactions will be released at lower salt concentrations. Proteins with stronger charges will have a higher affinity for the stationary phase and will remain bound to the column longer. In the chromatogram depicted above, protein B is released due a change in the conditions. The red line indicates some gradient applied to the system. When protein B is released, a second peak is created.

Regeneration
This step simply involves removing all bound protein from the stationary phase so that it is ready for another process.

Anion Exchangers
While the resin involved with an anion exchanger is positively charged, anion exchangers are named so because of their affinity to anions. To effectively bind proteins, the pH of the buffer in the system must be greater than the isoelectric point of the protein of interest, as proteins are negatively charged above their isoelectric point.

Anion exchangers can be classified as either weak or strong. The charged group on a weak exchanger is a weak base that easily loses its charge at a high pH due to deprotonation. On the other hand, strong anion exchangers are comprised of a charged group that is a strong base. Strong anion exchangers are able to maintain their positive charges across a variable pH range while weak anion exchangers tend to lose their charge as the pH increases. Examples of anion exchangers include the strong anion exchanger Q (quaternary resin), and the weak anion exchanger DEAE (diethylaminoethane).

Typically, the chromatography is performed using buffers at pH's between 7 and 10 and running a gradient from a solution containing just this buffer to a solution containing this buffer with 1M NaCl (Res1). The salt (in solution) competes for binding to the stationary phase, thus releasing the protein from its bound state. Proteins separate because the amount of salt needed to compete varies with the external charge of the protein. Anion exchange chromatography have been successfully used to clean up of a crude slurry, separation of proteins from each other, concentrating a protein, and the removal of negatively charged endotoxin from protein preparations (Res1).

Ion exchange in practice. In order to elute an unknown protein via ion exchange, a gradient of different concentrations of salt with appropriate buffer is required to test at which salt concentration our desired protein is eluted. In the case of anion exchange, DEAE is used to pack our column. A buffer, such as Bis-tris, used to equilibrate, wash and elute. A syringe pump is commonly used to pump various buffer solution via the column. Before an ion exchange, target protein is first equilibrated in Bis-tris buffer. Some protein might precipitate in Bis-tris hence solubility could be a potential problem during ion exchange. The column is first equilibrated with buffer with no salt. One salt commonly used is sodium chloride. Buffers that contain gradually increasing salt concentration is prepared, such as 50mM NaCl, 100mM NaCl, 150mM NaCl, etc. At last, a 1M NaCl buffer is used to completely clean up the column. All buffer needs to have their pH adjusted because protein are pH sensitive. As the diagram as suggested, fractions of increasing salt concentration are collected. After all fractions are collected, samples are taken to run an SDS page. With the gel, we can tell at which salt concentration our protein was eluted, and how well the separation is. The testing scale does not require a lot of protein mixture. In large scale protein purification, salt containing buffer can start rather high as an effective wash. Samples will be taken from all fractions to track our purification.

Cation Exchangers
Cation exchangers are named for their ability to attract cations or positively charged particles. In this case, the resin of the chromatography system is negatively charged and proteins will bind if the buffer pH is less than the protein’s unique isoelcetric point.

Just as with anion exchangers, cation exchangers can be classified as either weak or strong. A weak cation exchanger is comprised of a weak acid that gradually loses its charge as the pH decreases, while a strong cation exchanger is comprised of a strong acid that is able to sustain its charge over a wide pH range. Carboxymethal groups are used commonly as weak cation exchangers, while sulfopropyl groups are widely used as strong cation exchangers (Res1)

Designing a Cation Exchange Separation
Selecting the correct resin and buffer is critical as it determines the binding properties of proteins to the resin. In cation exchange, the protein of interests needs to be positively charge to bind to the negatively charged stationary phase. Suppose we have a mixture of the following protein :

 * At pH 4.8 cm, all protein will bind to the cation exchanger, with carbonic anhydrase binding the strongest to the stationary. If elution is performed at this pH, the protein will elute in the order of chymotrypsin, carboxypeptidase B, lysozyme, and carbonic anhydrase.

 * At pH 7.2 cm, carbonic anhydrase and carboxypeptidase B will elute in the wash (before the gradient is initiated), followed by chymotrypsin, then lysozyme during the salt gradient.

 * At pH 8 cm, only the lysozyme will bind to the stationary phase, the other three proteins will elute in the wash.

It is also possible to refine the salt concentration in solvents A and B so that the more weakly bound proteins will wash off the column in the wash, even if they are positively charged. For example, if you modify pH 4.8 cm to have 0.1 M NaCl in solvent A and 0.2 M NaCl in solvent B, you find that the most weakly bound protein (chymotrypsin) elutes in the wash, while the other three elute in the gradient.

Resources

 * 1) GE HealthCare Ion Exchange Animation
 * 2) Craig. P., Rochester Institute of Technology Ion Exchange Chromatography
 * 3) An online JAVA applet that simulates ion exchange chromatography can be found here: Ionex Simulation (Directions on how to set up experiments and run them using the simulation can be found at the link)
 * 4) Ion Exchange Image
 * 5) Bio-Rad Ion Exchange Chromatography & Products
 * 6) Sigma-Aldrich Ion-Exchange Ion-Exchange Resins and Related Polymeric Adsorbents
 * 7) Rohm and Haas Ion Exchange Resin Products


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Proteomics/Protein Separations - Chromatography/Anion exchangers

Anion Exchangers
While the resin involved with an anion exchanger is positively charged, anion exchangers are named so because of their affinity to anions. To effectively bind proteins, the pH of the buffer in the system must be greater than the isoelectric point of the protein of interest, as proteins are negatively charged above their isoelectric point.

Anion exchangers can be classified as either weak or strong. The charged group on a weak exchanger is a weak base that easily loses its charge at a high pH due to deprotonation. On the other hand, strong anion exchangers are comprised of a charged group that is a strong base. Strong anion exchangers are able to maintain their positive charges across a variable pH range while weak anion exchangers tend to lose their charge as the pH increases. Examples of anion exchangers include the strong anion exchanger Q (quaternary resin), and the weak anion exchanger DEAE (diethylaminoethane).

Typically, the chromatography is performed using buffers at pH's between 7 and 10 and running a gradient from a solution containing just this buffer to a solution containing this buffer with 1M NaCl (Res1). The salt (in solution) competes for binding to the stationary phase, thus releasing the protein from its bound state. Proteins separate because the amount of salt needed to compete varies with the external charge of the protein. Anion exchange chromatography have been successfully used to clean up of a crude slurry, separation of proteins from each other, concentrating a protein, and the removal of negatively charged endotoxin from protein preparations (Res1).

Resources

 * 1) Proteinchemist.com Ion Exchange Chromatography (IEC)
 * 2) Whatman Advanced Ion Exchange Cellulose and Column Chromatography

license: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/4.0/


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Proteomics/Protein Separations - Chromatography/Cation Exchangers

Cation Exchangers
Cation exchangers are named for their ability to attract cations or positively charged particles. In this case, the resin of the chromatography system is negatively charged and proteins will bind if the buffer pH is less than the protein’s unique isoelcetric point.

Just as with anion exchangers, cation exchangers can be classified as either weak or strong. A weak cation exchanger is comprised of a weak acid that gradually loses its charge as the pH decreases, while a strong anion exchanger is comprised of a strong acid that is able to sustain its charge over a wide pH range. Carboxymethal groups are used commonly as weak cation exchangers, while sulfopropyl groups are widely used as strong cation exchangers (Res1)

Designing a Cation Exchange Separation
Selecting the correct resin and buffer is critical as it determines the binding properties of proteins to the resin. In cation exchange, the protein of interests needs to be positively charge to bind to the negatively charged stationary phase. Suppose we have a mixture of the following protein :

 * At pH 4.8 cm all protein will bind to the cation exchanger, with carbonic anhydrase binding the strongest to the stationary. If elution is performed at this pH, the protein will elute in the order of chymotrypsin, carboxypeptidase B, lysozyme, and carbonic anhydrase.

 * At pH 7.2 cm, cm, carbonic anhydrase and carboxypeptidase B will elute in the wash (before the gradient is initiated), followed by chymotrypsin, then lysozyme during the salt gradient.

 * AtpH 8 cm, only the lysozyme will bind to the stationary phase, the other three proteins will elute in the wash.

It is also possible to refine the salt concentration in solvents A and B so that the more weakly bound proteins will wash off the column in the wash, even if they are positively charged. For example, if you modify pH 4.8 cm to have 0.1 M NaCl in solvent A and 0.2 M NaCl in solvent B, you find that the most weakly bound protein (chymotrypsin) elutes in the wash, while the other three elute in the gradient.

Resources

 * 1) Craig. P, Rochester Institute of Technology, Designing a Cation Exchange Separation


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Proteomics/Protein Separations - Chromatography/Molecular Exclusion

Chapter written by: Laura Grell and Alexander Butarbutar Contact llg3875@rit.edu or nbb3924@rit.edu for contributions Chapter modified by Kai Burnett and Dalia Ghoneim Contact kab9783@rit.edu or dxg6098@rit.edu

Molecular Exclusion

Molecular Exclusion / Gel Filtration / Gel Permeation / Size Exclusion is one of the simplest mechanism for separating molecules. This method separates proteins based on their size. The stationary phase is packed with porous bead with a range of pore sizes. When a mixture of protein is introduced into the column, a range of protein from small, intermediate and large will be filtered through the column
 * Small proteins can fit inside any pores and enter many pores before reaching the end of the column. Thus the movement of small proteins are greatly impeded which cause them to elute last in SEC.
 * Intermediate size protein may fit in some pores but not in all. These protein will generally elute somewhere between the small size and large size protein.
 * Large size proteins cannot enter the beads. Their movements in the column are not impeded by beads which make them the first protein to be eluded from the column.

In practice, gel filtration can be used to separate proteins by molecular weight at any point in a purification of a protein. It can also be used for buffer exchange - a protein dissolved in a sodium acetate buffer, pH 4.8, can be applied to a gel filtration column that has been equilibrated with tris buffer, pH 8.0. Using the tris buffer, pH 8.0, as the mobile phase, the protein moves into the tris mobile phase as it travels down the column, while the much smaller sodium acetate buffer molecules are totally included in the porous beads and travels much more slowly than the protein.

Resolution in Molecular Exclusion Chromatography
Achieving high resolution protein separation depends mainly on two factors:

 * 1) Sufficient selectivity: creating conditions that create enough space between sample zones
 * 2) Sufficient efficiency: counteracting "zone broadening effects"

The resolution can be expressed in the following formula: $$ R = \frac{Vr1 - Vr2}{(W1 + W2)/2}$$

Vr1 and Vr2: the elution volumes of two adjacent peaks. Vr2-Vr1 therefore represents the distance between two peaks.

W1 and W2: the widths of the peaks

Achieving Sufficient Selectivity
The numerator of the Resolution formula represents the selectivity or the distance between two different curves in a chromatogram. When the distance is too short, two adjacent curves may start to merge resulting in an adequate separation of the two proteins. The selectivity is controlled by two factors: the selectivity of the actual medium, and the length of the column.

Different mobile phase medium have different selectivity curves. Some mediums provide high selectivity while others provide lower selectivity. The medium chosen usually reflects the type and size of the proteins being separated.

The length of the column also has an effect on the selectivity. Molecular Exclusion does not involve any gradients; therefore, the molecules will move down the column at a fairly constant rate. If two molecules are traveling at different rates, increasing the travel distance will subsequently increase the distance between the two molecules. Increasing the column length is one method that can improve resolution. One caveat to this, however, is that as the column lengthens, the sample zones broaden. This zone broadening effect will counteract some of the gained resolution.

Achieving Sufficient Efficiency
Increasing the efficiency involves counteracting zone broadening effects. Ideally all of the sample of a specific protein should reach the end of a column at the same time; however, several factors have an effect on the level of zone broadening that occurs. The two most significant ones are: uniformity in beads, and bead size and flow rate.

It is essential to have uniformity in the beads in a column. Irregularities will can different parts of a sample to travel down the column at different speed. The range of time over which the sample reaches the eng of the column becomes large which is not ideal.

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Resources

 * 1) Craig, P., Rochester Institute Of Technology Gel Filtration Chromatography
 * 2) GE HealthCare Molecular Exclusion Animation || Introduction to Gel Filtration & Products
 * 3) Bio-Rad Size Exclusion Chromatography Kit || Size Exclusion Chromatography & Products
 * 4) Molecular Exclusion Image


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Proteomics/Protein Separations - Chromatography/Affinity

Chapter written by: Laura Grell and Alexander Butarbutar Contact llg3875@rit.edu or nbb3924@rit.edu for contributions Chapter modified by Kai Burnett and Dalia Ghoneim Contact kab9783@rit.edu or dxg6098@rit.edu

Affinity Chromatography
Affinity chromatography is a biochemical method designed to separate proteins from a mixed sample. It is one of the most commonly used techniques as it is very selective and effective at isolating proteins. The technique relies on unique interaction between a molecules with a ligand bounded to the matrix. Affinity chromatography have been used in a variety of applications, several types of matrix are readily available and can be purchased at various vendors. These matrices include interaction between:

Affinity chromatography has been successfully performed with a number of protein tags, one of the most common application is the use of poly-histidine tag. Extracting a protein from a gene product can be difficult task; one strategy that is commonly employed is to engineer a poly-histidine tag near the end of the gene. Typically, the histidine tag is relatively short and does not interfere with protein conformation. In affinity chromatography, the poly-histidine tag will bind to the nickel resin in the column.

Developing an effective affinity chromatography method involves:

 * 1) Finding a ligand that is specific enough
 * 2) Finding suitable conditions for binding between the target protein and the ligand as well as release the protein.

Stages in affinity chromatography
The affinity chromatography process can be separated into 3 main stages:

 * 1) Equilibration
 * 2) Application of Sample
 * 3) Elution

Equilibration
The first step in affinity chromatography is to prepare a stationary phase by immobilizing one of the two recognized components on an insoluble hydrophobic polymer such as agarose.

Application of Sample
A crude mixture containing the other recognized component is then applied to this stationary phase. Only those that are attracted to the stationary phase will bind to the column, the rest will simply be washed through the column. The success of this portion of the experiment is highly dependent on the target molecule's ability to sufficiently bind to the ligand. The equilibrium dissociation constant (KD) is a value that quantifies the strength of the bond between the target protein and its ligand. The lower the KD value, the stronger the bond. KD values between 10-4 and 10-6 are ideal for binding of the target molecule to the stationary phase.

If the KD value is too high, there will be leakage of the target protein. If the KD value is too low, the target protein will bind too strongly. This may result in "irreversible" adsorption to the stationary phase. Although in this case, the adsorption is not truly irreversible, the necessary elution conditions may be very harsh and may destroy the protein's ability to function.

During a wash step, which often includes a high concentration of salt to overcome weaker recognition interactions, all proteins except those that bind tightly to the stationary phase are eluted.

Elution

The wash is followed by an elution step in which a soluble form of the immobilized ligand is used to displace the bound protein from the stationary phase.

Since only the specific target sample can bind to the stationary phase, no fine-tuned elution gradient is necessary. In affinity chromatography it is possible to simply switch from binding conditions to release conditions immediately.

There are two possible ways to elute the target molecule from the stationary phase:

 * 1) Change the KD value from low to high. Ideal KD values for elution are usually between 10-1 and 10-2. The KD value can be altered by changing conditions such as the salt concentration, pH, temperature etc.
 * 2) Add a competitor that will displace the target protein from the ligand. This can involve adding free ligand that will bind to the target protein and wash out with the protein, or can be a competitor that binds to the ligand and in place of the target protein (Figures 2 and 3).

Target Molecules
There are three groups of target molecules that can be used in affinity chromatography:

 * 1) Specific binding based on Biological Activity: These include receptor binding sites, antibody binding sites,and enzyme active sites. These are used with their naturally occurring ligands or analogues of these ligands. It is possible to use these to separate groups of molecules if and analogue to a ligand has broad specificity.
 * 2) Prosthetic groups that occur in nature: An example of this is polysaccharides. This is usually only useful in performing group separations.
 * 3) Molecules engineered to contain an affinity tag: Examples of common tags are GST (Glutathione-S-Transferase) or proteins with accessible histidine residues. These affinity tags are almost always engineered into recombinant proteins for the purpose of separation.

Ligands

When using ligands that are mono-specific, it is difficult to find a commercially produced matrix specific for each case. Companies often sell "activated gels" that have active residues ready to be coupled to ligands. Ligands that are used in group-specific chromatography experiments are usually available commercially because they are more widely applicable.

Smaller ligands may result in some problems. One problem that may arise is steric interference. Another problem that may occur is blocked access to the ligand as illustrated in Figure 4. A spacer arm can solve this problem.

< Previous Page | Next Page >

Resources

 * 1) Davidson.edu Affinity Chromatography Methods
 * 2) GE Healthcare Affinity Chromatography Animation || Introduction to Affinity & Products
 * 3) Craig, P. Affinity Chromatography
 * 4) EYLaboratories, Inc - List of various ligand
 * 5) Affinity Image
 * 6) Bio-Rad Affinity Chromatography & Products


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Proteomics/Protein Separations - Chromatography/Immobilized Metal Ion Affinity Chromatography (IMAC)

Chapter written by: Laura Grell and Alexander Butarbutar Contact llg3875@rit.edu or nbb3924@rit.edu for contributions Chapter modified by Kai Burnett and Dalia Ghoneim Contact kab9783@rit.edu or dxg6098@rit.edu

Immobilized Metal Ion Affinity Chromatography (IMAC)
Immobilized metal ion/Metal Chelate affinity chromatography is a separation technique that is based on coordinate covalent binding between proteins and metal ions. Proteins have a wide variety of amino acids composition which, in effect, generates a range of different affinities towards metal ions. However, not many naturally occurring proteins have affinity for metal ions, so the technique is mainly used to purify recombinant proteins. For example, proteins can be engineered to contain a poly-histidine tail (histidine can generally act as a ligand towards divalent metal cations). If the stationary phase is immobilized with divalent metal cations, a mixture of proteins can be separated based on their ability to interact with the metal ions. Those proteins containing a higher number of histidine residues would be able to bind to the column more tightly than those with fewer histidine residues.

Several different types of immobilized metal ion column have been developed to separate various proteins (e.g. Fe, Co, Cd, Ni, or Zn). Protein separation in IMAC generally depends on the strength of the metal ion-protein bond. Thus, choosing the type of immobilized ion is crucial to successful protein separation. By far the most widely-used technique is to use an immobilized nickel column, and to engineer poly-histidine tags of six or more residues onto the recombinant proteins of interest. One thing to keep in mind is that the binding between metal ion and protein must be reversible, allowing elution of bounded protein at later steps. Three different elution strategies can be applied to IMAC competitive elution, stripping elution and pH Adjustment (Ref1).

< Previous Page | Next Page >

Resources

 * 1) Affiland Immobilized metal ion affinity chromatography (IMAC)
 * 2) GE Healthcare Immobilized metal ion affinity chromatography products
 * 3) Merck Metal Chelate Affinity Chromatography Using Tentacle Gels
 * 4) Bio-Rad Profinity IMAC Resins


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Proteomics/Protein Separations - Chromatography/Reversed Phase

Chapter written by: Laura Grell and Alexander Butarbutar Contact llg3875@rit.edu or nbb3924@rit.edu for contributions Chapter modified by Kai Burnett and Dalia Ghoneim Contact kab9783@rit.edu or dxg6098@rit.edu

Reversed Phase

Reverse Phase chromatography is a separation based on the solubility of the protein. The term ‘reverse’ was derived from its predecessor named ‘normal’ phase chromatography, which utilized a polar stationary phase such as silica. In reverse phase, the stationary phase is packed with silica modified with silyl ethers containing non-polar alkyl groups typically C8 or C18. This creates a hydrophobic stationary phase. The mobile phase on the other hand, contains relatively polar organic solvents such as methanol, butanol, isopropanol, acetonitrile (Ref1).

Utilization of these polar solvents introduces very harsh conditions for the protein, thus the method will generally work well for smaller and more stable proteins. All peptides and proteins carry a mix of hydrophilic and hydrophobic amino acids, but those with high net hydrophobicity will be able to participate in hydrophobic interactions with the stationary phase. As mixtures of proteins are applied to the column, polar proteins will elute first while non-polar proteins will bind to the column. Elution of the bound hydrophobic protein can be accomplished by increasing the concentration of organic solvent, which increases the retention time of a particular component. Reverse phase chromatography is commonly coupled with mass spectrometry in an effort to quantify the protein that is eluted from the column (Res1).

Stages in Reversed Phase Chromatography
The stages in Reversed Phase Chromatography are very similar to the stages found in Ion exchange Chromatography. Once again, the process can be summarized in four steps(Res2):

 * 1) Equilibration
 * 2) Sample Application
 * 3) Elution
 * 4) Regeneration

Equilibration
In this step the hydrophobic column is primed by applying the specific sample buffer. Because the column is hydrophobic, water molecules tend to be ordered at the junction between the column and the buffer.

Application of the Sample
In this step, the sample protein is injected into the system. Proteins in the mixture that have a high percentage of exposed hydrophobic amino acid residues will be adsorbed to the hydrophobic stationary phase. Other proteins in the mixture will be washed out.

Elution
This stage in the process involves changing the buffer conditions to elute the bound hydrophobic protein. The most common way to do this is to use a gradient that slowly increases the hydrophobicity using an organic buffer. The order in which proteins are desorbed is usually relative to the number of external hydrophobic residues each protein has (Res 3).

Regeneration
This stage involves washing off any remaining protein from the stationary phase and returning the conditions back to the way they were at the start of the process. This involves creating a less hydrophobic environment.

Example
Here is a sample diagram of a reversed phase separation run for 80 minutes, reaching a 70% organic solvent level with three elution peaks.

Note the gradient delay in this diagram. The delay is common to all liquid chromatography gradients, and is caused by a delay in the software telling the gradient to start pumping, and the solvent reaching the column. Also note the organic solvent concentration doesn't start at 0%. This is because the bonded hydrocarbons are much less efficient at binding to other hydrophobic molecules without a small amount of organic solvent present.

Optimization

In order to see greater peak separation, you can run the same proteins again for a shorter period of time, or lower the maximum amount of organic solvent. Using the above diagram, it appears that the last protein elutes at about 50% organic solvent concentration. After taking into account the delay, the last peak elutes and an adjusted 40% organic solvent concentration. A new separation can then be done, using a 40% maximum organic solvent concentration, which would generate much more widely separated peaks.

< Previous Page | Next Page >

Resources

 * 1) University of Adelaide Department of Chemistry Determination of caffeine in soft drinks by HPLC
 * 2) GE HealthCare - Reversed Phase Animation || Reversed Phase Products
 * 3) Andrew Guzzetta Reverse Phase HPLC Basics for LC/MS
 * 4) Proteinchemist.com Reversed Phase Chromatography (RPC)


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Proteomics/Protein Separations - Chromatography/Hydrophobic Interaction Chromatography (HIC)

Chapter written by: Laura Grell and Alexander Butarbutar Contact llg3875@rit.edu or nbb3924@rit.edu for contributions Chapter modified by Kai Burnett and Dalia Ghoneim Contact kab9783@rit.edu or dxg6098@rit.edu

Hydrophobic Interaction Chromatography (HIC)
Hydrophobic Interaction Chromatography is a separation technique that uses the properties of hydrophobicity to separate proteins from one another. In this type of chromatography, hydrophobic groups such as phenyl, octyl, or butyl, are attached to the stationary column. Proteins that pass through the column that have hydrophobic amino acid side chains on their surfaces are able to interact with and bind to the hydrophobic groups on the column.

HIC separations are often designed using the opposite conditions of those used in ion exchange chromatography. In this separation, a buffer with a high ionic strength, usually ammonium sulfate, is initially applied to the column. The salt in the buffer reduces the solvation of sample solutes thus as solvation decreases, hydrophobic regions that become exposed are adsorbed by the medium.

(Res1). The more hydrophobic the molecule, the less salt needed to promote binding (Res1). To elute the proteins, the salt concentration is gradually decreased, eluting bound species in order of increasing hydrophobicity. Additionally, elution can also be achieved through the use of mild organic modifiers or detergent (Res1).

The stationary phase is designed to form hydrophobic interactions with other molecules. These interactions are too weak in water. However, addition of salts to the buffer result in hydrophobic interactions. The following is a list of salts that increase hydrophobic interactions in the order of their ability to enhance interactions:

 * 1) Na2SO4
 * 2) K2SO4
 * 3) (NH4)2SO4
 * 4) NaCl
 * 5) NH4Cl
 * 6) NaBr
 * 7) NaSCN

Although reversed phase chromatography and hydrophobic interaction chromatography are very similar, the ligands in reversed phase chromatography are much more hydrophobic than the ligands in hydrophobic interaction chromatography. This enables hydrophobic interaction chromatography to make use of more moderate elution conditions, which do not disrupt the sample nearly as much.

< Previous Page | Next Page >

Resources

 * 1) Bio-Rad Hydrophobic Interaction & Products
 * 2) GE Healthcare Hydrophobic Interaction Chromatography Animation || Introduction to Hydrophobic Interaction & Products


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Proteomics/Protein Separations - Chromatography/Two Dimensional (2D) Chromatography

Two Dimensional (2D) Chromatography
Separation of highly complex mixture can be a very difficult task. The mixture can be distributed according to their molecular mass, chemical composition, functionality and architecture (Res 5). A single chromatography experiment may be inefficient in separating our proteins of interest. In 2D chromatography, different techniques are essentially combined to achieve a higher degree of separation. This can be done by an offline technique, where the result of one chromatography is injected manually into a second column chromatography or an on-line method, where the two columns are directly coupled through switches. (Ref 6).

2-D chromatography is considered a complement to 2-D electrophoresis. While 2-D electrophoresis separates proteins based on Isoelectric point and size, 2-D chromatography can use many different combinations of properties for separation. There are many different types of 2D chromatography, these include:

Multidimensional Protein Identification Technology (MudPIT) is a separation technique which utilized two chromatography techniques back to back and can be coupled to silica capillary (Res1). Typically the column consist of strong cation exchanger (SCX) followed by reversed phase (RP) material. One of the advantages of MudPIT is that band broadening are minimized and the capillary can easily be coupled directly with a mass spectrometer. The strategy initially involves denaturing the protein complex into small peptides. The samples are then loaded to strong cation exchanger (SCX) column subsequently followed by reversed phase column. A capillary can be set up to stream line the process with a mass spectrometer. 2D plot image from MudPIT
 * Multidimensional Protein Identification Technology (Mud-PIT)
 * Affinity-Reverse Phase
 * Size Exclusion Chromatography-Reverse Phase
 * Size Exclusion Chromatography-Ion Exchange Chromatography
 * Ion Exchange Chromatography-Reverse Phase

2-D chromatography is growing in popularity (Res 2) as an alternative to 2-D electrophoresis. It is good for separating proteins that have a more extreme range of pIs or a large range of molecular weights. A nice advantage of 2-D chromatography is that it can be coupled to a mass spectrometry device directly. In electrophoresis, the samples must be cut from the gel for retrieval.

An example of a setup utilizing 2D Chromatography can be seen below:

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Proteomics/Protein Separations- Electrophoresis/Introduction to Electrophoresis

Presentation
Introduction to Electrophoresis

Definitions
e•lec•tro•pho•re•sis (ĭ-lĕk'trō-fə-rē'sĭs) n. 1) The migration of charged colloidal particles or molecules through a solution under the influence of an applied electric field usually provided by immersed electrodes. Also called cataphoresis. 2) A method of separating substances, especially proteins, and analyzing molecular structure based on the rate of movement of each component in a colloidal suspension while under the influence of an electric field.

an•a•lyte (a-nə-līt) n. A chemical substance that is the subject of chemical analysis.

Electrophoresis Theory
Separation by electrophoresis depends on differences in the migration velocity of ions or solutes through a given medium in an electric field. The electrophoretic migration velocity of an analyte is:

$$v_p = \mu_p E$$

where E is the electric field strength and $$\mu_p$$ is the electrophoretic mobility.

The electrophoretic mobility is inversely proportional to frictional forces in the buffer, and directly proportional to the ionic charge of the analyte. The forces of friction against an analyte are dependent on the analyte's size and the viscosity (η) of the medium. Analytes with different frictional forces or different charges will separate from one another when they move through a buffer. At a given pH, the electrophoretic mobility of an analyte is:

$$\mu_p = \frac{z}{6\pi \eta r}$$ where r is the radius of the analyte and z is the net charge of the analyte.

Differences in the charge to size ratio of analytes causes differences in electrophoretic mobility. Small, highly charged analytes have greater mobility, whereas large, less charged analytes have lower mobility. Electrophoretic mobility is an indication of an analyte's migration velocity in a given medium. The net force acting on an analyte is the balance of two forces: the electrical force acting in favor of motion, and the frictional force acting against motion. These two forces remain steady during electrophoresis. Therefore, electrophoretic mobility is a constant for a given analyte under a given set of conditions.

Applications of Electrophoresis
Electrophoresis has a wide variety of applications in proteomics, forensics, molecular biology, genetics, biochemistry, and microbiology.

One of the most common uses of electrophoresis is to analyze differential expression of genes. Healthy and diseased cells can be identified by differences in the electrophoretic patterns of their proteins. Proteins themselves can also be characterized in this way, and some sense of their structure can be derived from the masses of fragments inside the gel.

There are many different types of electrophoresis, and each can be used for something different. Two-dimensional (2-D) electrophoresis, for example, has the ability to discern many more proteins than most of its contemporaries. Many of these methods will be discussed in detail throughout this chapter.

What Will Proteomics Gain from Microfluidic?
Proteomics contributes significantly to the discovery of proteins and their functions that influence the behaviors of an organism. Recent studies have focused on its roles to explore the proteins in a single cell and tissue level as they represent a fingerprint for each individual especially in terms of how a disease exhibits. Due to a limited amount of sample from a single cell or tissue, the study of proteome in these levels becomes a big challenge for a regular benchtop instrument. To facilitate this level of study, a microfluidic technology is introduced and developed into an essential tool for proteomics.

In addition to the capability to handle small sample, the microfluidic technology plays an important role in miniaturizing the entire system. As a result, a better performance in terms of less material consumption, faster processing time, more automated, and lower cost can be achieved. Another advantage in such microscale is that the mixing between the sample and reagents becomes more effective. A certain chemical process that usually takes hours can be completed in minutes. This benefit allows the microfluidic-based immunoassay to be used to monitor the progress of disease. And, with a reduction in cost, the microfluidic device becomes a perfect candidate for many point-of-care applications.

One of the most intriguing features provided by the microfluidic technology is its highly integrated capability with other systems especially with mass spectrometry. Multiplexing of various assays is another example. This multiplexing potential makes microfluidic-based devices a high throughput solution in (bio)chemistry and biomedicine.

Electrophoresis in Microfluidic Microsystems
Incorporating biotechnology with microfluidic makes a manipulation of very small volume of biological fluid not only feasible, but also effective especially by means of electrophoresis. Recent research and development efforts have been focused on inventing an electrophoretic microsystem that is fully automated, easy to customize for a specific need, and provides the results consistent with the gold standard. This microfluidic microsystem is usually referred to as a lab-on-a-chip. Among the microfluidic microsystems used in the analytical (bio)chemistry, the most widely used methods to control the transport of biomolecules or analytes are either gel or capillary electrophoresis.

Even though both techniques utilize the fact that biomolecules such as proteins, peptides, and DNA become charged in a buffer, the microfluidic gel electrophoresis operates differently from its capillary counterpart in terms of fluid dynamic. In microfluidic gel electrophoresis, the presence of porous gel medium prevents a bulk flow in a microfluidic channel. On the other hand, the bulk flow becomes an engineering factor that influences the electrokinetics of the analytes as the presence of electroosmotic flow needs to be considered as well in microfluidic capillary electrophoresis. In the analytical viewpoint, their electrophoretic separation methods are also different. The gel-based electrophoretic separation of biomolecules is based on the difference in size or molecular weight, while the capillary-based electrophoresis separation operates by taking the advantage of the difference in charge-to-mass ratio among biomolecules.

In comparison to the gold-standard methods, the analytical results obtained from the microfluidic gel and capillary electrophoresis are consistent with those from the traditional slab gel and capillary electrophoresis, respectively. In the design and application viewpoints, however, there are some advantages and disadvantages of these two methods needed to be considered. In terms of engineering design, the microfluidic gel electrophoretic system is much easier to customize for specific applications and multiplexing since no influence from the bulk flow needs to be considered. This makes it more straightforward to integrate extra features like sample preprocessing into the microfluidic gel electrophoresis. Nevertheless, since the gel-sieving medium is not required in the capillary-based microsystem, the reusability of the microfluidic capillary electrophoretic device is much higher than the gel-based counterpart. In terms of bio(analytical) applications, the microfluidic capillary electrophoresis has a drawback such that it operates poorly to analyze the charged particles of similar charge-to-mass ratios. It is worth noting that some microfluidic capillary electrophoretic system is capable of direct interface with mass spectrometry.

In this chapter, the device fabrication process, basic principle of operation, and some clinical applications of both microfluidic gel and capillary electrophoresis are described in detail. Even though this chapter is dedicated mainly to the microfluidic electrophoresis, the integration of additional features like sample preprocessing, detection, and quantification processes are also included. Unquestionably, this integration is made possible by the microfluidic technology.

Microfluidic Gel Electrophoresis
Utilizing microfluidic technology in gel electrophoresis provides several advantages to the study of proteome in many ways that cannot be achieved by the conventional methods. Faster processing time, more sensitive detection, more automated operation, and highly integrated system are the major benefits of using microfluidic. In addition, the microfluidic technology allows gel electrophoretic system to be easily customized for a specific application. For example, a microfluidic gel electrophoretic system can be designed such that off-chip processing can be eliminated. In fact, it can be integrated into the microfluidic-based system. The integration of sample preparation is one of the practical examples that not only simplify the experiment protocol, but also improve the detection sensitivity.

This section is dedicated to a customized microfluidic gel electrophoresis for immunoassay applications. The important aspects such as the fabrication process, principle of operation, and clinical applications will be discussed in detail.

I. Fabrication Process
Gel electrophoresis can be customized for a specific analytical study such as an immunoassay by using microfluidic technology. The customized fabrication process of microfluidic gel electrophoretic immunoassay to be described below is based on the device used by Herr et al. In their study, an antibody was used as a reporter to detect the presence of a particular protein as an antigen, potentially a disease biomarker. A step-by-step fabrication process is described as follows.

Step 1: Fabrication of Size-Exclusion Membrane

Materials :

 * 1) Degassed 22% (15.7:1) acrylamide/bis-acrylamide (6% bis-acrylamide cross-linker)
 * 2) 0.2% (w/v) VA-086 (photoinitiator)
 * 3) 1X Tris/glycine buffer

The first component of the microfluidic gel electrophoretic immunoassay to be fabricated is a size-exclusion membrane. This membrane is used to enhance analyte concentration, thus improving sensitivity of detection. A polyacrylamide gel is utilized to fabricate this membrane. It is designed such that the polyacrylamide gel has a pore size small enough for selectively allowing analyte molecules smaller than 10 kDa to pass through. The fabrication process begins with patterning a glass substrate to create microfluidic channels and chambers. This process is carried out by using regular photolithography and wet etch. Holes are drilled on the top glass cover. Both glass substrate and cover are then bonded together by anodic bonding. After the microfluidic structure is created, a solution of degassed 22% (15.7:1) acrylamide/bis-acrylamide (6% bis-acrylamide cross-linker) and 0.2% (w/v) VA-086 is introduced into the channel, as shown in the diagram. A syringe can be used to load the solution into the channel. The gel precursor solution is left for equilibration for approximately 30 minutes.

The size-exclusion membrane is fabricated using laser photo-polymerization. A 355-nm UV laser sheet is used to pattern the polyacrylamide gel at the specified location, shown in the diagram, to create the membrane profile. The polyacrylamide gel membrane is exposed to the laser until polymerized, which takes approximately 15 seconds. The remaining gel solution is vacuumed out, and the channels are then cleaned by rinsing with buffer.

Step 2: Fabrication of Separation Channel

Materials :

 * 1) Degassed 8% (37.5:1) acrylamide/bis-acrylamide (2.6% bis-acrylamide cross-linker)
 * 2) 0.2% (w/v) VA-086 (photoinitiator)
 * 3) 1X Tris/glycine buffer

After the size-exclusion membrane is fabricated, the next step is to construct a separation channel. This separation channel is the place where protein separation takes place. It contains a medium porosity polyacrylamide gel. Like the membrane, the separation gel is fabricated by photo-polymerization. To fabricate the separation channel, the separation gel precursor solution is carefully loaded into the microfluidic channel by a syringe. The gel solution is composed of the degassed 8% (37.5:1) acrylamide/bis-acrylamide (2.6% bis-acrylamide cross-linker), 0.2% (w/v) VA-086 photoinitiator, and 1X Tris/glycine buffer. The gel loading direction is indicated by the arrow shown in the diagram. The gel is loaded up to the specified location to define the separation channel.

Gel uniformity is a very important factor for the repeatability in the analysis. To guarantee uniformity, all microfluidic channels on the separation side of the membrane must be filled with gel before polymerization. Therefore, a gel plug is created to prevent the gel leakage during the subsequent gel loading. As shown in the diagram, the gel plug is fabricated by photo-polymerization such that the area not to be polymerized is protected by a dark-field mask. Usually, the photo-polymerization process takes about 10 minutes using a 100-Watt UV source.

Step 3: Fabrication of Loading Channel

Materials :

 * 1) Degassed 3.5% (37.5:1) acrylamide/bis-acrylamide (2.6% bis-acrylamide cross-linker)
 * 2) 0.2% (w/v) VA-086 (photoinitiator)
 * 3) 1X Tris/glycine buffer

In this step, the remaining microfluidic channels (without gel) on the separation side of the membrane are carefully filled with gel precursor solution as indicated by the arrow shown in the diagram. The solution contains the degassed 3.5% (37.5:1) acrylamide/bis-acrylamide (2.6% bis-acrylamide cross-linker), 0.2% (w/v) VA-086 photoinitiator, and 1X Tris/glycine buffer. This gel solution will generate a polyacrylamide gel with large pore size and define the loading channels.

After both separation and loading channels are filled with gel, the whole microfluidic device is exposed to UV for 15 minutes. As a result, the separation and loading gels are polymerized and define separation and loading channels, respectively. This microfluidic device is now ready to use.

Complete Microfluidic Device

The diagram shows the complete microfluidic gel electrophoretic device after fabrication. The device contains polyacrylamide gels with three different pore sizes. The gel with largest pore size is used in the loading channels to facilitate the electrophoresis of the sample and reagents by preventing bulk flow. The gel with intermediate pore size is used in the separation channel for protein separation. Finally, the gel with smallest pore size is used as a size-exclusion membrane for the enrichment process. The detail descriptions about their functions will be covered in the section entitled Principle of Operation.

In the device design, all the loading channels are connected to the loading areas, which are holes drilled at the beginning (before anodic bonding). The sample and reagents are loaded into there. In addition, some holes are used as reservoirs for waste collection. These are the places to which the electric current flows. It is worth noting that in addition to being the loading areas, they are also used as the insertion points for electrodes, which are connected to a programmable supply voltage source.

When not in use, this microfluidic gel electrophoresis device must keep submerging in the buffer and store at 4°C.

II. Principle of Operation
Based on the same principle of electrokinetic transport of charged molecules, the microfluidic gel electrophoresis works similarly to the regular slab gel electrophoresis, but with much faster processing, more sensitive detection, and highly automated-integrated systems. In this section, the fundamental operation of the microfluidic gel electrophoresis will be discussed using the microfluidic structure described in the previous section.

The microfluidic gel electrophoretic system to be discussed consists of a structure containing microfluidic channels/reservoirs, power supply, and fluorescence detection system. The channels and reservoirs used to transport the analyte are designed such that they are filled with large pore size polyacrylamide gel. This large pore size gel facilitates the electrokinetic transport of the analyte by preventing the bulk flow. The power supply is designed so that it is programmable. A pair of electrodes and their polarities can be assigned instantaneously. This provides a much better control over the electrokinetic process only attainable in microfluidic environment. Also integrated into the system is the fluorescence detection capability. It is used to detect the analyte of interest and quantify its concentration.

In this section, a step-by-step operation procedure including the principle behind each step will be discussed, assuming that the target analytes are negatively charged. Note that the discussion will mainly focus on the immunoassay application, which is more generalized to the study by Herr et al. A little modification can be made to the system to be used in other applications.

Loading a Reporter of Known Concentration

As a tool for immunoassay study, a reporter specific to a particular protein of interest is used. Acting like a receptor with high affinity for a target protein, an antibody can be used as the reporter for the detection of protein or antigen being investigated. Without loss of generality, the term reporter will be used throughout this section.

To begin using the device, the microfluidic channels need to be filled with buffer solution. Then, the reporter solution of known concentration together with sample solution is loaded into the designated reservoir. Typically for the microfluidic device, the volume required per analysis is roughly in the order of a few tens of microliters. For the purpose of detection and quantification, the reporter is usually labeled with fluorescence tag. The detection and quantification methods will be discussed in detail later.

After the fluorescently labeled reporter is loaded, a pair of electrodes connected to the reporter and sample waste reservoirs are activated as shown in the diagram. The electric potential is then applied across both electrodes. As the reporter molecules become charged in the buffer, they are electrokinetically moved toward the sample waste reservoir as indicated by the red arrows. The reporter molecules, however, are blocked by the size-exclusion membrane, which allows only particles of size less than 10 kDa to pass through. In this loading step, only the ionic buffer can penetrate the membrane. At the end of this first electrophoresis, the increasing number of the fluorescently labeled reporter molecules gathers at the membrane on the gel side as shown in the inset.

Loading a Sample Containing Analyte of Interest

The next process is to load the sample electrokinetically to combine with the reporter, which already gathered at the membrane. Like the reporter, the analyte molecules become charged in the buffer. To begin the electrophoresis, the electrode contacted to the reporter reservoir is deactivated. The electrode contacted to the sample reservoir is switched on instead. The potential across these two electrodes causes the charged molecules of proteins in the sample to move electrokinetically toward the sample waste reservoir as indicated by red arrows.

Once the charged molecules of the analyte arrive the membrane, some of them whose size less than 10 kDa pass through the membrane and are collected in the sample waste reservoir. Only the larger molecules including the protein of interest stay on the gel size of the membrane as shown in the inset. This process helps increase the concentration of the analyte, improving the probability of binding between the reporter and target protein. However, the non-target proteins become more concentrated as well and might cause a reduction in signal-to-noise ratio level. Using the reporter of high binding specificity can alleviate this problem. It is worth noting that the sensitivity and dynamic range of the microfluidic gel electrophoretic immunoassay depend on this enrichment process.

It needs to be designed carefully so that the electrophoresis occurs long enough so that more reporters and the target proteins are bound together. Normally, the concentration of the reporter used is much higher than that of the target protein. As a result, all the target protein molecules would be more likely to be detected by the reporter molecules. At the end of this second electrophoresis, the reporter and its complex including the remaining non-target proteins stay on the gel side of the membrane.

Avoid Running Electric Current Through Membrane During Separation

It has been reported that the gel electrophoretic separation experienced irreproducible results when running the electric current through the membrane. Therefore, it is necessary to avoid applying the electric potential across the membrane especially during the separation process. One possible solution is to program the power supply such that the electrodes are switched before the target analytes enter the separation gel. The diagram shows the switching process that satisfies this requirement.

According to the diagram, the electric potential is initially applied across the membrane just to transport the analytes away from the membrane. Before the analytes enter the separation gel, the electrode at the upper left buffer reservoir is deactivated. At the same time, a new electrode with the same polarity at the upper right buffer reservoir is turned on. It can be seen that the flow of negatively charged analytes remains in the same course toward the buffer waste reservoir.

It is worth noting that there are other combinations of electrodes that can be used to perform this step as well. The concept is simply not to allow electrophoresis to occur across the membrane during the separation step. At the end of the third electrophoresis, all charged molecules are ready for the gel electrophoretic separation.

Gel Separation

In this step, the reporter molecules and their complex are separated by means of gel electrophoresis. The process continues by electrokinetically carrying the charged molecules into the separation gel. Since the reporter molecules and their complex have similar charge-to-mass ratio, the separation between these two species is based only on the difference in their molecular weights (MWs). It is worth noting that all non-target proteins are not investigated in this immunoassay. Only the reporter molecules and their complex are under examination.

Since the reporter molecules are labeled with fluorescent dye, a single-point laser can be used to induce the fluorescence. This laser-induced fluorescence (LIF) is monitored by a detector that will detect the presence of the unbound reporter and its complex, and then relate the intensity of the detected fluorescence to the concentration of both species. This fluorescence detection system is placed across the separation channel near the buffer waste reservoir. The concepts of detection and quantification are described next.

Fluorescence Detection and Results

The single-point laser-induced fluorescence (LIF) is a mean used to detect and quantify the reporter molecules and their complex. When the molecules across the laser beam, the attached dyes emit fluorescence whose intensity is measured by the detector. The detector then generates the electropherogram corresponding to the measured intensity. The area under the electropherogram peak can be related to the concentration of the target protein.

The unbound reporter can be distinguished from the complex based on the fact that the unbound reporter molecule has lower molecular weight (MW) than its complex. Therefore, the unbound reporter molecules travel electrokinetically faster in the separation gel and are detected first. The corresponding intensity gives the first peak in the electropherogram.

In case of a single pair of ligand and receptor, there will normally be two peaks in the electropherogram. Usually the first peak belongs to the receptor or reporter of smaller MW, followed by the second peak corresponding to the ligand-receptor complex of larger MW. This information can also be represented by gel-like plots, which are computer-generated. In the gel-like plots, the topmost band represents the unbound reporter molecules that arrive first. The brightness of the band conveys the information about the degree of fluorescence intensity detected. The width of the band corresponds to the width of the peak, which conveys the information about the migrating time of the analyte.

It is worth mentioning that the probability of binding between both receptor and ligand is a significant factor to the sensitivity and accuracy of the immunoassay. Therefore, the enrichment process plays an important role in raising this probability. To enhance this enrichment process, either preprocessing the sample (off-chip) or increasing the period of the electrophoretic sample loading is proven to be useful.

III. Clinical Applications
One of clinical applications reported was the use of this microfluidic gel electrophoretic device in assisting oral diagnosis. It was used for early diagnosis of the periodontal disease and to monitor the disease state and its development from human saliva. Periodontal disease or periodontitis is a putative oral disease that destroys collagen and causes a major tissue damage, connective tissue attachment loss, and bone loss. Found in the periodontitis patient saliva were matrix metalloproteinase-8 (MMP-8), interleukin-1 beta (IL-1B), and C-telopeptide pyridinoline cross-links (ICTP). These proteins were identified as disease biomarkers and became the target proteins for detection and monitoring this oral disease. Herr et al. demonstrated the detection and quantification of MMP-8 using this microfluidic gel electrophoretic device.

In their study, the monoclonal antibody for MMP-8 was used as a reporter, which had the specificity for binding only to the MMP-8 in the saliva. The use of this monoclonal antibody ($$a$$MMP-8) had an advantage in that it eliminated the need for a matched pair antigen-antibody. In the reporter mixture, the bovine serum albumin (BSA) protein standard was also added as a reference. Both $$a$$MMP-8* and BSA* were fluorescently labeled for the detection purpose. Note that the asterisk is used to denote the fluorescently labeled analyte. According to their study, 1 nM of $$a$$MMP-8* and 1 nM of BSA* were used in the mixture.

In the microfluidic device, $$a$$MMP-8* bound only to MMP-8 biomarker forming a complex of similar charge-to-mass ratio. The unbounded antibody and its complex were separated from each other by the separation gel based on the difference in their sizes and detected individually as described in the Principle of Operation for microfluidic gel electrophoresis. In this immunoassay application, the BSA* with the highest mobility was detected first, followed by the remaining $$a$$MMP-8* and MMP-8 complex, respectively. The corresponding peaks were shown in the electropherogram and gel-like plots. The peak area or the width of the band in gel-like plot was used to calculate the concentration of MMP-8.

To be able to quantify the concentration of the endogenous MMP-8 in the saliva sample collected from the patients, a calibration curve was required to be generated first. The calibration curve was obtained from the analyses of known MMP-8 concentrations. A series of experiments were performed by adding known concentrations of the recombinant MMP-8 in the diluted saliva from healthy patients and then running the microfluidic gel electrophoresis. From the electropherogram, the peak areas of the MMP-8 complex were normalized with the peak area of BSA* and then plotted against the corresponding concentrations. A nonlinear least-squares fitting method using a four-parameter logistic model was used to obtained the calibration curve. Based on this calibration curve, the concentration of the endogenous MMP-8 complex was predicted from its normalized peak area. It was reported that the average concentrations of the MMP-8 in the healthy and disease subjects were 64.6 +/- 16.4 ng/mL and 623.8 +/- 204.0 ng/mL, respectively. It is worth noting that the concentration of MMP-8 in the patients classified as periodontally diseased exhibited the dynamic activity of the disease.

The validity of the results obtained from the microfluidic gel electrophoresis was verified by comparing with those obtained from the conventional enzyme-linked immunosorbent assay (ELISA). As reported by Herr et al., the results obtained from the microfluidic device were highly correlated with those from ELISA with r2 = 0.979, where r is Pearson product-moment correlation coefficient. This microfluidic-based immunoassay, however, had several advantages over the conventional counterpart in that it bypassed the time-consuming reaction and washing steps required by ELISA, was more automated, required only single antibody making it applicable for wider range of applications, and needed much less amount of saliva sample per analysis. In addition, it did not require surface immobilization of the antibody.

In addition to measuring the concentration of MMP-8 in the periodontitis patients, a clinical examination was also performed to correlate the analytical data with physiological symptoms. It was found that MMP-8 was highly correlated with bone and tissue loss, but having no correlation with bleeding upon probing. With highly sensitive detection of MMP-8 biomarker, this microfluidic-based immunoassay could provide early diagnosis that would improve the clinical treatment of this disease. Furthermore, with the use of MMP-8 inhibitor to reduce collagen degradation, it was promising that this microfluidic gel electrophoretic device could be used to monitor and track the progress of MMP-8 inhibitor therapy as well.

Microfluidic Capillary Electrophoresis
In this section, another type of microfluidic electrophoresis will be described. Unlike microfluidic gel electrophoresis, microfluidic capillary electrophoresis operates on the principle of electrokinetic of bulk flow. The difference in charge-to-mass ratio of analytes is the fundamental of electrophoretic separation in capillary microfluidic channel. Without the need of sieving medium, the microfluidic capillary electrophoresis becomes more favorable than its gel counterpart in terms of the ease in fabrication and reusability. The detail fabrication process and principle of operation including clinical applications of this device are described as follows. Note that the fabrication process and principle of operation explained below are based on the published work by Backofen et al. 2002.

I. Fabrication Process
The fabrication of microfluidic structure using PDMS is easier and much less expensive than using glass. The process begins with creating a master for molding the PDMS, which can be obtained by patterning a photoresist. After developing, a mixture of PDMS and cross-linker is poured over the patterned photoresist and then cured. The PDMS is peeled out and cut to create an access to the reservoir. In the final step, this patterned PDMS is attached permanently to a glass base and ready to use. The detail step-by-step procedure is described below.

Creating a Master for PDMS Molding

Materials :

 * 1) Photomask
 * 2) 4-inch silicon wafer
 * 3) SU-8 50 negative photoresist
 * 4) Propylene glycol methyl ether acetate developer

Beginning with a preparation process, a photomask is created first. The photomask used is usually a contact photomask that can be made of glass or simply a transparency with the designed microfluidic structural layout printed on it. The photomask is used to transfer this microfluidic pattern on to a photoresist, which is spin-coated onto a silicon wafer. It is worth noting that the choice of photoresist to be used affects the design of photomask. For example, if using the negative photoresist, all the microfluidic patterns need to be designed using clear field so that the UV light can go through and polymerize the contact areas of the negative photoresist. The unexposed areas will be washed away in a developer. On the other hand, the designed patterns are the dark field of photomask for the positive photoresist.

Before coating the photoresist, the silicon wafer is needed to be cleaned first. A standard RCA cleaning process is applicable. Then, the negative photoresist is spin-coated on the silicon wafer and pre-baked. Exposing under the UV light, the coated negative photoresist is patterned with the designed microfluidic layout from the photomask. The exposed photoresist is then post-baked and developed. As a result, only the exposed areas of the negative photoresist remain as shown in the diagram.

Molding PDMS to Fabricate Microfluidic Structure

Materials :

 * 1) PDMS oligomer
 * 2) Sylgard 184 cross-linker

After the master for PDMS molding is ready, a mixture of PDMS and cross-linker is prepared. A 10:1 ratio of the polymer and cross-linker is applicable. The degassed mixture is then poured onto the wafer to cover the entire photoresist master. Finally, the molded PDMS is cured in the oven at 70 °C for 1 hour. This process step is summarized in the diagram. In addition, a cross-sectional view of the PDMS molded on the wafer is also provided.

After curing, the PDMS is peeled out of the wafer. The PDMS with the transferred microfluidic structure is further processed by creating an opening for each circular reservoir region. This opening will be used to load sample or reagent and a place to apply vacuum and pressure, which is regulated by a syringe.

Finalize Microfluidic Capillary Electrophoretic Device

Materials :

 * 1) Glass plate
 * 2) Needle probe

A glass plate with coated patterned electrodes is used to enclose the microfluidic channels and reservoirs, and at the same time, provides electrical connections to the fluid in reservoirs. The electrodes can be fabricated on glass substrate by means of evaporation of chrome and platinum. Chrome is used as an adhesive layer between glass substrate and platinum electrodes. The patterning of the metals can be achieved by a regular photolithography, which requires another photomask (a transparency), to form the electrodes on glass. It is worth noting that a combination of patterned electrodes and needle probe is used in the final microfluidic capillary electrophoretic system as shown in the diagram. The platinum needle probe is used to supply high electric potential for electrophoretic purpose and simultaneously as an electrochemical sensor.

The PDMS is permanently attached to the glass plate by oxidation using plasma. A covalent bonding between oxygen and silicon atoms (O-Si-O) provides a permanent seal between both materials. The complete microfluidic capillary electrophoretic device is shown in the diagram.

II. Principle of Operation
In general, the operation of microfluidic capillary electrophoretic device can be described by three fundamental steps – loading sample/reagents, forming sample plug, and electrophoretic separation. The loading process described here involves manual loading using syringes to regulate the flow. After all sample and regents are loaded, a sample plug is formed. This step needs to be designed carefully since the concentration of the analytes relies on this step. In the last step, the analytes are separated electrophoretically. Unlike the size-based separation in the (microfluidic) gel electrophoresis, the separation by (microfluidic) capillary electrophoresis is based on the differences in charge-to-mass ratios of the analytes.

It is worth noting that the microfluidic capillary electrophoretic system being discussed here is used as an analytical platform where all the sample preprocessing steps are performed off-chip. The following description focuses mainly on the analysis of negatively charged analyte in the sample. Note that the principle of operation described below is based in part on the published work by Backofen et al. 2002.

Loading Process

The system setup begins with loading the microfluidic reservoirs with a buffer and preprocessed sample solution such that all the reservoirs except the sample reservoir contain the buffer. Continuing with filling the microfluidic channels with buffer, syringes are connected to the top openings of the reservoirs and used to regulate the flow. Vacuum and pressure are applied until the channels are filled with the buffer.

The next process is to equilibrate the sample and buffer by means of electrophoresis. To do so, the high voltage sources are connected to the system as shown in the diagram. One possible configuration is the use of multiple supply units such as two units of U1 = 0.5 kV and a single unit of U2 = 1.2 kV. This process allows the buffer and sample to localize in the channels as shown in the diagram.

Forming Sample Plug

For the sample to be analyzed, a predefined and controllable portion of the sample needs to be injected into the separation channel. This predefined and controllable portion of the sample is referred to as a sample plug. According to the microfluidic structure being discussed, the sample plug is formed by a hydrodynamic flow of the sample solution due to the difference in the solution levels in the reservoirs. This hydrodynamic flow only occurs when all the high voltage sources are switched off.

As depicted by the diagram, the portion of sample solution flows through the small connecting channel into the channels leading to the buffer and buffer waste reservoirs. This sample portion can be divided into two ports indicated by blue and yellow dashed outlines as shown in the inset. The blue outline is the sample plug whose volume can be specified by design. The yellow dashed outline indicates the portion of the sample that will flow back toward the sample waste reservoir when switching the high voltage sources back on.

Electrophoretic Separation

When switching on all the high voltage sources, the negatively charged analyte molecules and buffer ions start to flow again by electrophoresis. At the small connecting channel, the portion of sample solution begins to split into two parts and then move away in the opposite directions. As depicted in the inset, the part of sample solution indicated by blue-dashed outline, or the sample plug, electrokinetically moves along the separation channel toward the buffer waste reservoir. On the other hand, the part of sample solution outlined by the yellow dashed line electrokinetically move away toward the sample waste reservoir. The small connecting channel is then filled by the buffer as occurred originally.

In the separation channel, the negatively charge analytes in the sample plug are separated due to the differences in their charge-to-mass ratios. Each analyte is detected by the electrochemical sensor placed at the end buffer waste reservoir. For the electrochemical sensor used in this microfluidic capillary electrophoretic system, the detection of each analyte is based on a reduction in voltage drop across the needle probe and reference electrode. These voltage drops can be plotted in the electropherogram and can be used to study the analyte components such as proteins or peptides in the sample solution.

III. Clinical Applications
The detection and quantification of biomarkers in patients with skin lesions reported by Guzman et al. 2008 is one of the clinical applications for the microfluidic capillary electrophoresis. Based on the same fundamental concept described previously, a more sophisticated design of microfluidic capillary electrophoresis so-called the immunoaffinity capillary electrophoresis (IACE) was used in their study. IACE is a lab-on-a-chip that utilizes the microfluidic technology to incorporate the affinity-based purification, enrichment, and electrophoretic separation processes in one single microchip.

In the clinical study of biomarkers for this inflammatory disease, IACE was used to analyze the micro-dissected samples from the patients with different stages of skin damage. Twelve different antibodies were used to capture twelve corresponding target proteins/peptides being considered as biomarkers. This affinity-binding also assisted the isolation of target analytes from non-target analytes in microfluidic environment, thus improving the subsequent enrichment process and reducing background noise during the analysis. As reported, the integrated enrichment feature increased the sensitivity and enhanced the low detection limit of IACE. The concentration as low as a few nanogram per milliliter could be detected and quantified. With this highly sensitive capability, it was also reported that IACE could be used to monitor the progressive patterns of the disease. The results obtained by IACE were proven consistent with the gold standard like the traditional histopathology.


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User:Businessdepot

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Proteomics/Protein Separations- Electrophoresis/Databases

 * World-2DPAGE, listing, query portal
 * This site contains a listing to other 2D PAGE Database servers by organism, as well as a Web portal to query simultaneously World-Wide proteomics databases.

 * SWISS-2DPAGE,
 * This database contains data on proteins identified on a large collection of 2-D PAGE and SDS-PAGE reference maps. The database is linked to many other proteomics ressources. It has a graphical interface, as well as a search engine offering many various queries.

 * Proteome 2D-PAGE Database,
 * 2D-page protein files for various bacterial proteins can be searched by organism or by clicking on a spot on a reference map.

 * BIOBASE Biological Databases: Proteome Database,
 * The site contain databases for human, S.cerevsiae, S.pombe, C. elegans,and pathogenic fungi. Search by name of protein. Links to NCBI Webpage

 * Danish Centre for translational breast cancer research,
 * Human 2D-page protein files can be searched by name, keyword, organelle or by clicking on a spot on a reference map. This site also includes a 2D gel immunoblot gallery, which displays whole proteins reacted with antibodies.

 * Gelbank,

 * BPP-2DPAGE,
 * Hematopoietic cell lines can be searched by clicking a spot on a reference map.

 * Parasite Host Cell Interaction 2D-PAGE database,
 * Parasite host cell interactions can be searched by author, description or by clicking a spot on a reference map.

 * Yeast Protein Map,
 * This site contains 2D gel maps of yeast proteins listed by gene and ORF names.

 * SIENA-2DPAGE,
 * This database can be searched by description, author or by clicking on spots.

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Proteomics/Protein Separations- Electrophoresis/Web Pages

 * American Electrophoresis Society,
 * This website has descriptions of current electrophoretic separation and detection methods
 * Corbett Learning,
 * This site includes a simulation of the basic steps involved in electrophoresis
 * Separations Now,
 * This site includes new articles concerning recent advances in electrophoresis
 * ExPasy Proteomics Server,
 * This site is monitored by the Swiss Institute of Bioinformatics. ExPasy stands for Expert Protein Analysis System.  Databases, proteomics tools and information on molecular biology are provided.
 * Joint Proteomics Laboratory,
 * This is the webpage of the Ludwig Institute for Cancer Research whose focus is on developing techniques for protein separation.

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Proteomics/Protein Separations- Electrophoresis/Further Readings

Open Source Articles

 * Discussion of the strengths and weakness of using 2-D gel electrophoresis coupled with mass spectrometry for proteomics research
 * Beranova-Giorgianni, S. (2003)  Proteome analysis by two-dimensional gel electrophoresis and mass spectrometry:  strengths and limitations.   Trends in Analytical Chemistry 22(5), 273-281.[]

 * Discuss the capabilities of 2DGE when combined with mass spectrometry in proteome analysis
 * Stephen J Fey and Peter Mose Larsen(2001) 2D or not 2D: Current Opinion in Chemical Biology 2001, 5:26–33[]

 * Discusses a variant of QPNC-PAGE
 * Kastenholz, B. 2004. Preparative native continuous polyacrylamide gel electrophoresis (PNC-PAGE): An efficient method for isolating cadmium cofactors in biological systems. Anal. Lett. 37, 657-665.

 * Contributions of QPNC-PAGE
 * Kastenholz, B. 2006. Important contributions of a new quantitative preparative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE) procedure for elucidating metal cofactor metabolisms in protein-misfolding diseases – a theory. Protein Pept. Lett. 13, 503-508. Special issue [Hot Topic: Recent Advances in Protein and Peptide Chemistry Related to Cellular Regulation (Guest Editor: John W. Ho)].

Subscription Required Articles

 * Review of high-throughput separation methods including a discussion on gel-electrophoresis based techniques
 * Suresh, S., Mohan, S., Mishra, G., Hanumanthu, G., Suresh, M., Reddy, R., Pandey, A. (2005)  Proteomic resources:  Integrating biomedical information in humans.  Gene 364, 13-18.  Abstact[]

 * Discuss the merits of 2D-PAGE when coupled with ICAT in quantitative proteome analysis
 * Marcus Smolka‡, Huilin Zhou§, and Ruedi Aebersold§(2002) Quantitative Protein Profiling Using Two-dimensional Gel electrophoresis,Isotope-coded Affinity Tag Labeling, and Mass Spectrometry. Molecular & Cellular Proteomics 1:19–29,[]

 * Discuss the merits and drawbacks of 2DGE and ICAT
 * Wayne F Patton*, Birte Schulenberg and Thomas H Steinberg(2002) Two-dimensional gel electrophoresis; better than a poke in the ICAT?. Current Opinion in Biotechnology, 13:321–328 Abstract []

 * Highlights the advances of 2D gels in quantitative proteomics
 * Kathryn S Lilley, Azam Razzaq and Paul Dupree(2001)Two-dimensional gel electrophoresis: recent advances in sample preparation, detection and quantitation. Current Opinion in Chemical Biology, 6:46–50 Abstact[]

 * Review of historical and current multi-dimensional peptide separation methods
 * Issaq, H., Chan, K., Janini, G., Conrads, T., Veenstra, T. (2005)  Multidimensional separation of peptides for effective proteomic analysis.  Journal of Chromatography B 817 (1), 35-47.

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Proteomics/Protein Separations - Chromatography/Ultra Performace Liquid Chromatography

Ultra Performance Liquid Chromatography
High performance liquid chromatography has proven itself to very useful in many scientific fields, yet forces scientists to consistently choose between speed and resolution. Ultra performance liquid chromatography (UPLC) eliminates the need to choose and creates a highly efficient method that is primarily based on small particle separations.

Waters ACQUITY Ultra Performance LC systems have been developed to take into account all the advantages that small particle separations currently have over HPLC. Many of these advantages are primarily based on the theories behind liquid chromatography. In general, increasing the efficiency of a separation will also increase its resolution. Since both efficiency and optimum flow rate are inversely proportional to particle size, a decrease in the particle size will increase efficiency and speed up the flow rate.

In the ACQUITY system, the particle size is decreased to 1.7um compared to 3.5um or 5um. The particles were specifically designed to withstand wide ranges of pressure and pH, have a high load capacity, and improve efficiency. Other innovations to the chromatography method include a high pressure solvent delivery system, to take into account the smaller particle size, fast injection cycle sample management, and specialized detectors with fiber optic flow cell design.

Resources

 * 1) Waters - Ultra Performance Liquid Chromatography
 * 2) Waters - UPLC New Boundaries For The Chromatography Laboratory


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Proteomics/Proteomics and Drug Discovery/Structure-Based Drug Design

 * 1) Introduction
 * 2) Structure Based Drug Design
 * 3) Virtual Compound Libraries
 * 4) Docking and Scoring
 * 5) Software Tools
 * 6) Protein Aggregation

Overview
In the early 1980s, researchers were not able to take advantage of structure-based methods in the drug discovery process. This was due to a number of factors, the most important of which were a lack of computing power and docking programs that could test potential models as well as a lack of interest in the established community due in part to the aforementioned lack of tools. However, in the 1990s, computing power and available programs increased exponentially as well as the ability to obtain cheaper, more reliable x-ray crystallographic structures necessary for any type of computational study. This marked the start of a new era and the first attempts and successes were published, with the two most famous being the HIV-1 protease inhibitors and renin inhibitors (to combat hypertension) (Lunney). In contemporary drug discovery, structure-based methods are an integral part of the drug development process. This change can be attributed to the rapid advances in genomics and structural biology, as well as developments in information technology. The advancements of technology in several fields that are vital to the drug discovery process increased the pace of drug development. However, many years of research are still required until a drug which is both effective and tolerated by the human body is marketable (Anderson). Despite this new technology and increased funding by pharmaceutical companies, the issues with finding safe, effective compounds have resulted in there being no significant increase in the rate of therapeutic agent release to the general public. (Lindsay)

The drug discovery process includes many steps. First, the drug target has to be selected. In most cases this will be a protein; however, recent studies have shown that RNA, with its well defined secondary-structure, is also an effective drug target. DNA is also a target of newer drugs, especially in chemotherapy (see below). At least 25 percent of currently marketed drugs target G-protein-coupled receptors (GPCRs), while another 22 percent affect the function of ion channels, proteases, kinases, and nuclear hormone receptors. Only 2 percent of therapeutic targets in 2003 were DNA or RNA. Once a target is selected, it has to be cloned and purified in order to determine the structure. The most common method used for structure determination is X-ray crystallography, but NMR, which when applied to proteins uses multidimensional experiments that increase unique signals and the signal to noise ratio, and homology modeling, which uses closely related protein structures to clarify the structure of an unclear protein, are often employed as well. New methods in phase determination of molecules, along with complete automation, have allowed for high-throughput X-ray crystallography; this has greatly enhanced the speed of structure determination. As soon as the structure of the target is known and a potential ligand binding site is identified, computational methods can be used to dock a large number of small molecules into the specified position. These small molecules, which are stored in a database, are then scored and ranked based on their steric and electrostatic (see also, ionic bonds) interactions with the target site. The best performing compounds, called ‘hits’, are selected for biochemical assays and further testing. The hit compounds have to be effective at very low concentrations, at least at the micromolar level. The hits are further optimized into the more potent ‘leads’ by chemical synthesis to improve their potency. Selected leads are further scrutinized by rigorous cytotoxicity tests, pharmacokinetic studies, and toxicology investigations to search for side effects followed by the eventual phase 0, I, and II clinical trials for successful drug prototypes (Alanine, Anderson).

Drug Target Selection and Identification of the Target Site
The selection of the drug target is mainly based on biological and biochemical considerations. Proteomics as a tool in this area is still relatively limited due to the sheer complexity of protein expression in any given cell. Despite this, proteomics’ usefulness has grown in other areas of the drug discovery process including biomarker identification and tracking. The ideal drug target for structure-based drug design should bind a small molecule and should be closely associated with the disease. The small molecule then either changes the function of the drug target, or in case of a pathogenic organism, inhibits the function of the target. This will ideally lead to the cell death of the pathogen. In the latter case, the drug target should only be present in the diseased cells or pathogen and should have a unique function that allows for and encourages this selectivity. Furthermore, the uniqueness of the drug target guarantees that another pathway cannot restore the function of an inhibited target. The structure-based search for anti-cancer and autoimmune drugs is much more challenging, since the drug targets regulate essential cell functions. Hence, these targets are not unique and isolated; inhibition of their function not only affects mutant or over/under-activated cells, but also normal cells. For example, the phosphoinositide 3-kinase (PI3K) pathway is both involved in cellular growth and has been directly linked with pancreatic cancer activation (Reddy). Trying to separate convoluted pathways such as this make the drug discovery process much more complex than with other diseases. Another option for targeting in cancer is DNA. Cisplatin and Bleomycin cross-link and cleave DNA respectively and are used to slow down cell division, particularly in cancerous tissues. (Singh)

The ligand binding site of a drug target should be a pocket containing both hydrogen donors and acceptors, as well as hydrophobic residues. In many cases, the selected target location is the active site of an enzyme, as with sildenafil citrate (Viagra), which targets the catalytic subunit of NADPH (Jeremy); however, it can also be an assembly or regulatory site, as is the case with the phosphotransferase regulatory domain the Bacillus subtilis Spo0f protein, a histidine kinase (Dai-Fu). Even protein-protein interaction sites, which are often large and planar, have been selected as target sites (2-oxoglutarate, a naturally occurring molecule, affects the monomer-monomer interactions of GlnK, an ammonia transporter [AcrB] inhibitor (Anderson, Stroud).

Proteomics as a Tool to Discover Biomarkers
‘Biomarker’ is short for ‘biological marker.’ A biomarker is a molecule, indicator, or test that can be used to measure such processes as disease progression, infection stage, and drug efficacy, as well as other various biological functions. They can also be used as part of safety studies for therapeutic agents. Although the most common biomarkers in use today are small molecules and proteins, the growing fields of pharmacogenetics and pharmacogenomics are attempting to utilize genotypes, haplotypes, and single nucleotide polymorphisms as biomarkers (Frank).

With the growing emphasis on biomarkers as indicators, the National Institutes of Health (NIH) began a ‘Biomarkers and Surrogate Endpoint Working Group.’ This organization has set up a classification system for biomarkers. Type 0 biomarkers are more symptomatic in nature and track disease progression over its complete history. They are used in phase 0 clinical studies. These clinical studies use well developed assay techniques in highly regulated populations for specific durations. The goal of these studies is to achieve simple positive or negative results for the drug or system being studied. Type 1 biomarkers are used to track any type of compound injected into the biological system. The most familiar aspects are drug trials, where the researchers are looking for a specific effect. The effect observed may be positive or negative. The final classification is Type II biomarkers. These biomarkers are used to determine a ‘surrogate endpoint.’ A surrogate endpoint, as defined by the FDA, “is a marker – a laboratory measurement or physical sign – that is used in the therapeutic trials as a substitute for a clinically meaningful endpoint that is a direct measure of how a patient feels, functions or survives and is expected to predict the effect of the therapy.”  In other words, a surrogate endpoint moves beyond the concept of a single biomarker and into the realm where many or no biomarkers may be sufficient. Other symptoms including overall health and mortality may be studied to determine the efficacy of a treatment regimen. Although surrogate endpoints are still in the early stages, two examples that have been accepted are blood pressure and cholesterol, which have a firm connection to cardiovascular health and mortality (Frank).

In addition to the above effects, a biomarker should correlate well with the disease condition and minimize the number of false positives, as well as false negatives. Thus, a biomarker should be able to accurately discriminate between a normal and a infected condition with high reproducibility. The challenge in proteomics research is to identify unique biomarkers from complex biological mixtures that unequivocally correlate with the disease condition. Biomarkers can be utilized for many purposes. Also, established biomarkers are useful as risk factor indicators, capable of providing information to show that a person is susceptible to a disease. QT prolongation, a measure of change in the ventricular electrical cycle, is used as a assessment of a patient’s chances for survival after heart attack, as is troponin T, a cardiac enzyme whose levels rise following a heart attack. 5-hydroxytryptophan is a metabolic precursor to seratonin, and has been found to localize around neoplasms in neuroendocrine tissue. Labelling of these molecules allows for visualization of these events by fluorodeoxyglucose positron emission tomography (FDG-PET), which is used in visualizing many types of malignancies. Another PET application uses SPA-RQ, a neurokinin-1 binder, whose injection is used to track binding of the drug Aprepitant™, a drug used in chemotherapy patients to control vomiting (Frank).

There are currently many more biomarkers being studied and in development, and this number will continue to grow as our understanding of the body evolves. The combination of tracer molecules and imaging techniques, as in 5-hydroxytryptophan, creates a powerful method for visualizing areas that should not be touched surgically except as a last resort. The growing use of proteomics to find biomarkers is important as well. Recent advances in analytical techniques such as mass spectrometry and column chromatography have allowed for more comprehensive studies of protein expression. In addition, 2-dimensional electrophoresis provides an overall view of expression, in similar fashion to gene studies. Proteomics studies using these techniques have identified ovarian cancer, macular degeneration, and lipoprotein composition. Development of these techniques will undoubtedly increase in the future (Frank).


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Proteomics/Proteomics and Drug Discovery/Contacts

Contacts
Anthony Esposito (age5719-AT-rit-DOT-edu)

Lukas Habegger (lh9357-AT-rit-DOT-edu)


text: Proteomics/Proteomics and Drug Discovery/References