• Packing the column
• Coupling ligand amines to agarose beads
  
• Coupling ligand carboxyl groups to agarose beads
  
• Magnetic bioseparation techniques
• Magnetic bead recovery techniques

1. Washing the beads : BioScience Agarose Beads are supplied fully hydrated and contain either 0.03% sodium azide or 17% ethanol as a preservative.
   
   
   1. In either case, the gel beads shold be placed in at least 3 bead volumes of distilled, deionized, or reverse-osmosis (RO) treated water and gently stirred for 10 minutes.
      
      
   2. The bead slurry should then be passed through a filter which will retain the beads.  A sintered glass filter in a Buchner funnel attached to a vacuum flask is quickest.   Note : do not overdry the beads.  If the the surface of the beads begins turning an opaque white, you are beginning to dehydrate the bead surface and low concentration (< 4%) beads can irreversibly dehydrate under these conditions.
      
      
2. Equilibration in starting buffer :  Place the filtered beads in 3 bead volumes of the desired application buffer and stir gently for about 10 minutes.
   
   
3. Degassing the bead/ buffer slurry : Place the beads slurry back in the vacuum filter flask and stopper the top before applying a vaccuum for about 5 minutes. Occasional swirling of the container will help to fully degas the slurry. Note do not use a stir bar in preference to manual swirling. ( save the degassed buffer for a later step).
   
   
4. Filling the column with buffer : Fill the column about 1/3rd full with degassed, starting buffer and dislodge any air bubbles that might be trapped in the column end pieces or bed support membrane.  Use a combination of tapping the column and draining the buffer through the exit stopcock to accomplish this.  Close the stopcock and replenish the degassed to 1/3rd the column height.
   
   
5. Filling the column with beads :  Fill the column with a filling funnel and reservoir and add the 50:50 bead/buffer slurry ( a convenient pouring concentration if well mixed).  Allow the beads to settle until ther is a bed height of about 4 cm.
   
   
   1. Open the stopcock and allow buffer to flow out until a stable , well-packed bed of beads has formed.
      
      
6. Connecting the column to the pump: Connect the top of the column or flow adapter ( recommended for SEC, IEC, HIC but is less critical for affinity chromatography, AIC) to the pump or Marriotte flask using some appropriate, non-leaching plastic tubing.  Fill the tubing and pump in the standard manner to insure that they are free of air bubbles.
   
   
   1. If a column reservoir is used, drain any excess buffer from it and attach the flow adapter to the column.
      
      
   2. Pump or drain at least 4-5 bead bed volumes of buffer though the column at a slightly higher flow rate than will be used during the planned chromatography.
      
      
7. Adjusting for bead bed volume changes :  Turn off the pump and close the column outlet.  Reposition the flow adapter until it is just making contact with the top of the gel bed.
   
   
8. Final equilibration with starting buffer ;  After the gel bed had equilibrated for about 10 minutes with the starting buffer, the column is ready for sample application.
   
   
9. Sample application :  The liquid sample is applied as a thin, uniform layer to the top of the gel bed and the buffer flow ( either by gravity, using a Marriotte flask or using a pump) is begun.
   
   
10. Chromatography :  Buffer flow is continued until the desired separation of sample components has been achieved and each component has both eluted from the column and been collected.  Quantitation of each component can be done either continuously, during elution, or after collection.
    
    

A) Coupling an Amine-containing Ligand to the Aldehyde Moiety of a Glyoxylated Agarose Bead:

The aldehyde groups of Glyoxal Agarose Beads react with ligands containing primary amine groups to form Schiff Base Intermediates, which are selectively reduced by cyanoborohydride. Sodium borohydride should not be used for this procedure because it reduces both the unreacted aldehyde and Schiff Base groups thereby lowering maximum coupling capacity. The higher the pH of the buffer used in the reaction, the greater the rate and extent of Schiff Base formation and ligand coupling. The resultant ligands formed are much more stable and less prone to leaching than those coupled using CNBr (cyanogen bromide).

Procedure:

1. Determine the amount of ligand you want to immobilize
   1. The amount of ligand required will depend on how much "target" substance you want to isolate in one use of the affinity beads to be prepared.
   2. Estimate the binding ratio of ligand to target substance (or assume it's 1:1)
   3. From (a) and (b) above, you now know the number of µmoles or µ equivalents of ligand to be bound to the beads.
   4. Glyoxal agarose beads contain at least 20µ equivalents of aldhyde groups per ml of beads. Calculate the volume of beads required to bind the desired amount of ligand.
2. Preparing the Coupling Buffer
   1. Using the Sigma Coupling Buffer (C 4187): This is highly recommended since this ready-to-use buffer minimizes the need to inventory the concentrated solutions of cyanoborohydride.
   2. Preparing your own coupling buffer: Follow the steps outlined at the end of this procedure in "Stock Solution Preparation".
3. Preparing the Glyoxal Agarose Beads: Filter a slurry of the glyoxal beads on a sintered glass filter or equivalent and weigh out the amount of beads required (1d). Equilibrate the beads in 3 volumes (1g. beads = 1ml) of coupling buffer for 15 minutes.
4. Equilibrating the beads: Filter the beads and add them to 2 bead volumes of fresh coupling buffer. Stir for 15 minutes.
5. Add the ligand as a dilute solution: Dissolve the ligand in water, saline, or coupling buffer and add it to the beads with stirring. Note: no amine-containing buffers should be present in the ligand soulution.
6. Allow coupling reaction to go to completion: Allow the reaction to proceed for at least 2 hr followed by filtering the beads and resuspension in 2 volumes of coupling buffer.
7. Block unreacted glyoxal sites: Mix 20 µl of ethanolamine for every ml of glyoxal beads with 5 ml of water and add the resultant solution to the ligand-coupled bead slurry and allow the reaction to proceed for 1 hr. Then wash the beads on the filter funnel with at least 5 volumes of water or buffer (but not coupling buffer).
8. Storage: The ligand-coupled glyoxal beads should be stored in a preservative-containing buffer which is suitable for the ligand.

Coupling Buffer Stock Preparation:

1. Alkaline buffer: 0.2 M disodium phosphate ( Sigma # S-9290) or 0.2 M sodium borate (Sigma # S-9640) solutions should be prepared. Dissolve the required amount of either salt in water. Note: The sodium borate is available as the sodium tetraborate decahydrate ( 76.2 g/L) as well as the tetrahydrate or metaborate salts.
2. Cyanoborohydride stock solutions: Prepare a 2 M solution of sodium cyano-borohydride - in the hood. Dissolve 6.3 g of sodium cyanoborohydride in water. It is best to let this solution stand overnight before use to decompose any residual sodium borohydride which might be present.
   Re: Safety:
   1. Cyanoborohydride solutions should NEVER BE ACIDIFIED ( i.e. pH reduced below 7.O).
   2. Perspecitve : Cyanogen bromide solutions have been used routinely for more than 40 years to produce cyanogen bromide activated agarose beads. All of the same safety precautions necessary for CNBr activation also apply for cyanoborohydride reduction.
   3. Always use unbreakable containers to minimize the likelihood of spills.
3. Coupling Buffer Preparation: Add 1 ml of cyanoborohydride stock solution from (B) above to either alkaline buffer solution (A) to form the coupling buffer. Because this buffer is more alkaline than the Sigma coupling buffer ( #1a above), it maximizes both the rate and extent of intermediate Schiff Base formation.

References:

1. Shainoff, J.R., "Zonal Immobilization of Proteins ", Biochem and Biophys. Res. Commun. 1980, 95, 690.
2. Shainoff, J.R. "Glyoxal Agarose" , U.S. Patent 4,275,196.
3. Guisan, J.M et. al, "Immobiliztion of enzymes on glyoxal agarose", Methods in Biotechnology, 277-287 ( 1997).
4. Hermanson, G.T. et. al, "Immobilized affinity ligand techniques", pg. 69-75, Academic Press, Inc. , San Diego, CA (1992).
5. Hearn, M et. al, J. Chromatog. 185, 453 (1979).
6. Lane, C.F., "Sodium Cyanoborohydride- a highly selective reducing agent for organic
   functional groups ", Synthesis, 3, 135-146 ( 1975).
7. Mosbach, Klaus, in "Methods in Enzymology" , Vol. XLIV: Immobilized Enzymes, 1976
   Academic Press, NY.
   

B) Coupling a Carboxyl-containing Ligand to the Amine Moiety of an Aminoethylated Agarose Bead:

Procedure:

1. Wash the required amount of aminoethyl (AE) agarose beads in 3-4 bead volumes of
   0.5 M NaCL.
2. For every 25 ml of AE agarose beads, prepare 1- 1.5 mM solution of EEDQ in 25 ml of ethanol. Use water instead of ethanol for the water soluble CDi's .
3. Add 0.5-1.5 mM of the ligand in 50% ethanolic EEDQ to the AE beads. The ligand can be dissolved in water for water-soluble CDi's . The volume of the ligand solution should not exceed 1.0 ml/ 2.5 ml of AE beads to avoid excess dilution.
4. Agitate preferable by rocking or swirling. Note: a typical magnetic stir bar or overhead
   stirrer-impeller will tend to damage the gel beads - particularly at high speed. Stir or mix overnight at room temperature ( 20-25°C).
5. Wash the AE beads with 50% ethanol for EEDQ, otherwise with water for other CDi's, until excess reagents or byproducts are eliminated.
6. Repeat washing with 3-4 bead volumes of 1 M NaCL followed by a rinse with DI or DO water.
7. The ligand is now coupled to the AE beads via an amide bond and the beads may be either used for affinity chromatography or stored in 0.03% sodium azide ( 4-10°C).

Coupling Reagent Nomenclature and Acronyms:

1. A water soluble carbodiimide (CDi) is preferable:
   1. CMCi : 1-cyclohexyl-3(2-morpholinoethyl) carbodiimide-metho-p-toluene sulfonate
   2. or EDCi: 1-ethyl-3(3-dimethylaminopropyl) carbodiimide hydrochloride.
2. An Alchohol/water soluble CDi is next best: EEDQ : (N-ethoxycarbonyl-2-ethoxy-1,2-
   dihydroquinoline)
3. Less desirable, unless hydrophobic solvents are used: DCC : dicyclohexyl carbodiimide.
   
   

1. A chosen volume of agarose beads are washed with distilled water (> 4 bead volumes) and drained- but not dehydrated-on a sintered glass Buchner funnel attached to a vacuum filter flask. It is particularly important that any alcohols or glycol preservatives be completely removed from the beads at this stage, since they would otherwise compete with agarose hydroxyls (for CNBr) and result in lower levels of agarose activation.
   
   Re: Agarose bead concentration: the beads can have an agarose concentration between 1 and 10 % agarose.
   
   
2. The washed beads should be free of any supernatant water but still moist and not  beginning to dehydrate - particularly if they are < 4 % in concentration since dehydration can irreversibly  change bead porosity.
   
   Note : Steps 3 - 8 & 12 MUST BE DONE IN THE HOOD.  CNBr has the capability to release poisonous cyanide gas IF ACIDIFIED. AS A RESULT, IT IS IMPORTANT TO KEEP THE PH OF THE CNBr REACTION IN THE ALKALINE pH RANGE ( ideally pH 10.2 - 11.0).  Keeping the reaction mixture cold is also important to obtaining maximum activation and avoiding premature hydrolysis of activated hydroxyl groups.
   
   
3. The beads are then slurried with an equal volume of cold (20°C) distilled water and gently stirred in the hood.
   
   
4. To control pH, a solution of NaOH should be prepared.  The amount of NaOH required depends on the amount of CNBr added.  For 5- 10 ml of agarose beads(and 1- 3 g CNBr), a 2 M solution should be prepared.  Note that the NaOH neutralizes the HBr formed when CN reacts with a hydroxyl group. For larger quantities of agarose beads (100- 200 ml and 20-30 g.of CNBr), a higher concentration of NaOH ( like 6M or 8M ) helps to minimize the total volume of the reaction mixture.
   
   
5. The beads are then activated with CNBr by adding the required amount directly to the cold,stirred bead slurry( 50- 200 mg of CNBr/ml of beads; depending on level of activation desired.
   
   
6. NaOH solution is IMMEDIATELY added so that the pH is adjusted to between pH 10.5 and pH 11 for the remainder of the reaction.  As the activation proceeds the pH will be seen to fall and more NaOH solution will need to be added.
   
   Note : Cyanogen bromide (CNBr) is available from the Aldrich Division of Sigma-Aldrich( cat# C91492-25 g. for the 25 g. quantity; other quantities available).
   
   
7. The reaction is usually complete in 6 - 12 minutes ( i.e. when the pH has stopped decreasing and no more NaOH is needed to keep it in the pH 10.5- 11.2 range.
   
   
8. The activated beads are then washed with 10- 15 bead volumes of cold(10-20°C) 0.1 M NaHCO3 on a sintered glass Buchner funnel.
   
   
9. The resultant agarose beads are now ready for coupling to a suitable protein ligand (i.e having at least one primary amine).
   
   
   1. The CNBr-activated beads are suspended in one volume of cold (10-20°C)0.1 M NaHCO3 and gently stirred.
      
      
   2. An appropriate amount of the protein to be coupled is also dissolved in one volume of  0.1 M NaHCO3 and then added to the beads with continued stirring.
      
      Note: There should be at least a 30 % excess of the protein (i.e milliequivalents of amine)to be coupled than calculated CNBr-activated sites ( in milliequivalents). For maximum coupling capacity, using as much as 20 times the desired amount of coupling ligand produces the best results.
      
      
10. The ligand coupling step is allowed to proceed with continued stirring overnight at 5-10 °C.
    
    
11. On a sintered glass funnel, the beads are then washed free of excess protein (i.e. ligand)and the excess ligand recovered as needed.   The beads are washed sequentially with  2 bead volumes of 0.5 M NaCl, 0.1 M borate buffer, 1 M NaCl, 0.5M NaCl, 0.1 M NaOAc, 1 M NaCl, and finally with 0.5 M NaCl.
    
    
12. Blocking any unreacted sites : The ligand-coupled beads are then resuspended in 2 bead volumes of 1 M NaCl,stirred gently, followed by addition of a 2:1 mole ratio of ethanolamine to CNBr used.  The ethanolamine reaction is allowed to proceed for 1-2 hr.
    
    
13. Final wash & storage:  the beads should be washed with several bead volumes of 0.1 M saline until there is no odor of residual ethanolamine. The beads should be placed in a suitable preservative solution ( 0.03 % sodium azide or equivalent) and stored under refrigeration ( 4-10°C) until used.
    
    Note : The beads should NOT be frozen since non-crosslinked beads will be irreversibly damaged by freezing.

Reference(s) :

1. Z. Er-el, Biochem.Biophys. Res. Commun., 49, No.2, 383-386 (1972).
   
   
2. R. Axen and S. Ernback, Eur. J. Biochem. 18, 351 ( 1971).
   
   
3. J. Porath, Nature(London) 218, 834 (1968).

All of the important parameters for effecting a successful magnetic bioseparation are discussed below followed by a step-by-step protocol.  The low cost, speed and simplicity of this protocol compared to a comparable affinity column technique is dramatic.

A. The Magnetic Bead :

1. Shape :  It is important that the bead be spherical.  Irregular shaped beads will
   tend to clump during recovery and redisperse poorly during subsequent steps.
   
   
2. No Non-Specific Bonding :  If the composition of the bead or method of ligand attachment produces ionic or hydrophobic groups - these will result in the binding of substances  other than those having an affinity for the  ligand.  The highly porous agarose gel found in BioScience Magnetic Beads,  is an ideal neutral polysaccharide having no ionic groups or  hydrophobic groups and also readily allows for convenient ligand immobilization .
   
   
3. Ligand Binding Capacity vs. magnetic composition:  The bead must have sufficient magnetite to be attracted by a suitable magnet but excess magnetite will reduce the potential binding capacity per unit volume of beads.  At the other extreme, if some beads in a population do not have sufficient magnetite, they will tend to be lost during wash steps.
   
   
4. Buoyancy:  The magnetic beads should have good suspension characteristics in water so they can be conveniently stirred with just a stir bar or overhead stirrer but do not require expensive special equipment.
   
   

B) The Magnet :

1. Type : A permanent magnet is highly preferred to avoid loss of magnetism over time or the need to remagnetize the magnet.
   
   
2. Strength :  A magnet having a 5 lb. lift capacity is optimal for recovering Magnetic BioScience Agarose beads.  The strength of the magnetic field produced by any magnet decreases as the inverse square of the distance from the magnet.
   
   
3. Composition: Neodymium magnets are preferable to most alternatives.  These magnets are made from a combination of metals and rare earth minerals which make them both strong and compact.  This type of magnet is supplied free with qualifying orders of Magnetic BioScience Beads.
   
   
4. Shape : The magnet shape can be adjusted to the method and scale of use. Rod, disk, plate or doughnut shapes are all common.
   
   
5. Size:  As with shape, above, the size of the magnet can be scaled to the recovery method used.
   
   
6. Safety :  Magnets can damage magnetic media like : floppy disks, credit cards, magnetic I.D. cards, cassette tapes, video tapes, and similar products.  They can also damage televisions, VCR’s computer monitors and other CRT electronics. As a result, a magnet should never be placed near any of these products.  A Neodymium Magnet should never be brought near a person with a pacemaker or similar medical device because it could alter it’s operation. Neodymium Magnets should not be either machined or burned and will lose their magnetic properties if heated above 80°C.

The photo sequence below demonstrates two ways, among many, in which a magnetic bioseparation can be accomplished.

Technique #1 ( for small vessels):

1. The magnetic agarose beads ( ~7 ml) having an attached ligand, are added to a graduated 50 ml plastic test tube and allowed to settle by gravity to measure their settled bead volume.  
   
   
   
   
2. A solution containing the target substance for the ligand (i.e. “ligate”) is added to the tube and the beads are shaken for several minutes - or whatever time is required for optimal binding between the ligate and ligand.
   
   
   
   
3. A magnet is placed at the side of the test tube and in 1-2 seconds the magnetic beads will be seen to cluster- securely- along one side of the tube.  The supernatant solution- now free of ligate- can be poured off.  
   
   
   
   
4. The tube is filled with a suitable wash solution or buffer and the tube is reshaken to resuspend the magnetic beads.  After a minute, the magnet is again used to immobilize the beads at the side or bottom of the tube, allowing for the supernatant to be decanted -while the magnet is held firmly in place.  A 2nd or 3rd such wash can quickly be done as needed.
   
   
5. The tube is then filled with a solution which is designed to break the bond between the ligand and ligate, thereby freeing the latter into solution. The solution chosen is the same one that would be used in a comparable affinity separation which is done in a column.
   
   
6. The  magnet is placed at the side of the test , as described in steps 1.3 & 1.4, above, and the supernatant - CONTAINING THE RECOVERED LIGATE - is carefully collected and the magnetic beads are washed in water followed by storage (4-10°C) in a suitable storage solution such as 0.03 % sodium azide.
   
   

Technique # 2 ( for large volumes of dilute ligate):

1. A Neodymium Magnet ( 0.5”diameter x 0.25”thick) having a 5 lb. lift capability, is attached to a standard stir-bar retriever.  
   
   
   
   Note: A typical stir-bar retriever does NOT have sufficient magnetic force to achieve good recovery of magnetic beads.  After attachment of a small, Neodymium Magnet to its tip, however, it works nicely.
   
   
2. A sufficient amount of ligand-attached  magnetic agarose beads are added to the solution containing a dilute concentration of ligate.  The volume of beads used is calculated based on the binding capacity of the beads in relation to the expected amount of ligate present in the solution to be harvested.
   
   
   1. The beads are gently stirred (overhead stirrer) or agitated ( roller bottle or equivalent).
      
      
   2. Note : Do Not use a magnetic stirrer or magnetic stir bar in an attempt to stir the magnetic beads - BECAUSE THEY WILL CLUMP AROUND THE STIRBAR AND MAGNETIC STIRRER AND NOT MIX PROPERLY.
      
      
3. Sterilizing a sheet of Saran® or similar plastic film :  In this procedure, the film acts as a removable magnet cover which allows the beads to be recovered quickly from a large volume of solution by moving the stir bar retriever/ magnet assembly through the solution - thereby increasing the local magnetic field on the beads by bringing the magnet to them instead of expecting them to come to the magnet .  This result is rapid recovery without the huge magnet that would otherwise be needed to attract all  the magnetic beads contained in a large vessel.
   
   
   1. A sufficiently large sheet of Saran® Wrap or similar plastic film should be wrapped around the stir-bar retriever and Neodymium Magnet on it’s tip, such that it  extends above the maximum point to which it will be immersed.   The Saran® is then secured above the maximum point of immersion.
      
      
   2. The Saran®-wrapped stirbar retriever/magnet should then be placed in boiling water until sterilization is assurred. Alternate methods of sterilization can be used.
      
      
   3. The Saran®-coated stirbar retriever can then be immersed in the test tube  or swept through a much larger volume and the magnetic beads are recovered in seconds.
      
      
      
      
      1. The magnetic agarose beads remain tightly bound to the Saran-coated stirbar retriever when the magnet is removed.
         
         
         
         
      2. The magnetically adhered beads can then be swirled in wash solution(s) followed by being swirled in a solution designed to break the bond  between ligate and ligand -as with technique #1.
         
         
4. The Saran-coated stirbar retriever is then placed in a sterile container and the film can then be peeled from the magnet such that the beads are either deposited in the sterile container or remain adhered to the outer Saran® surface - where they can be easily washed off with a gentle stream of water or buffer.
   
   
   
   Note: It requires virtually no force to peel the magnetic agarose beads away from the Neodymium Magnet at the tip of the stirbar retriever.
   
   
5. The magnetic agarose beads can then be treated with a solution designed to break the bond between ligand and ligate - so the target substance can be recovered.
   
   
   
   
6. The magnetic agarose beads can be recovered for future use by storing them in a suitable, refrigerated preservative ( 0.03% sodium azide).
   
   

Technique #3 :

1. Simply place a suitable flat magnet underneath a flat-bottomed vessel containing the magnetic agarose bead suspension.
   
   
2. In a short while, depending on the height of the vessel, the magnetic agarose beads will settle to the bottom- due to both the force of gravity and the additional magnetic force.
   
   Note: The larger the magnetic beads, the faster will be their rate of settling.