have contributed to the poorer-than-expected retention for y-globulin and hemoglobin, since in these cases the field and the flow would act in opposite directions. Zone Spreading. Theoretical plate data for hemoglobin in the acetate buffer are shown in Figure 6. These data confirm the theoretically predicted trend: greater field strength, and consequently greater retention, leads to a greater number of theoretical plates and therefore a smaller plate height. A quantitative comparison with theory, however, was not regarded as very meaningful because of the geometrical disturbances in the channel mentioned earlier. If one assumes a rigid channel, plate heights are roughly double the theoretical values. Separations. The efficacy of separation is determined by relative retention and by the extent of zone broadening. Figure 7 shows separations of albumin and hemoglobin in the pH 4.5 acetate buffer at field strengths increasing from 2.95 V/cm to 4.43 V/cm. Clearly, increased field strength magnifies differences in R and leads to improved separation. A similar result has been observed in sedimentation FFF (20). Figure 8 shows a series of separations of albumin and yglobulin in the pH 4.5 acetate buffer. While the improvement in separation from E = 1.97 V/cm to E = 2.95 V/cm results from increasing differences in R , the improvement seen from E = 2.95 V/cm to E = 3.44 V/cm is mainly due to a slight narrowing of the zones. This is not unexpected ( 9 ) ,as noted in the theory section, but extraneous factors may be involved as well. Separations were often clouded by an erratic baseline. The best possible baseline slope was used in constructing Figures 7 and 8, but this correction was not adequate to bring all peaks back to baseline; thus the peaks appear to stop short.
CONCLUSIONS In summary, the flexible channel column achieves reasonable separation. Moreover, the separations can be controlled
by the judicious variation of parameters. The theory serves to explain the principal trends. On the negative side, some disturbing anomalies make quantitative predictions of retention difficult. Flow parameters are also difficult to control. It is clear that room exists for improvement of the present system. It may be, also, that some alternative system will prove more tractable and provide better agreement with theory.
LITERATURE CITED (1) "Electrophoresis", M. Bier, Ed., Academic Press, New York, 1959. (2) K. D. Caldwell, L. F. Kesner, M. N. Myers, and J. C. Giddings, Science, 176, 296 (1972). (3) J. C. Giddings, Sep. Sci., 1, 123 (1966). (4) J. C. Giddings, J. Chem. Educ., 50, 667 (1973). (5) E. Grushka, K. D. Caldwell, M. N. Myers, and J. C. Giddings, in "Separation and PurificationMethods", Vol. 2, E. S. Perry, C. J. Van Oss, and E. Grushka, Ed., Dekker, New York, 1974, p 127. (6) M. E. Hovingh, G. H. Thompson, and J. C. Giddings, Anal. Chem., 42, 195 (1970). (7) J. C. Giddings, F. J. F. Yang, and M. N. Myers, Sep. Sci., I O , 133 (1975). (8) J. C. Giddings, J. Chem. Phys., 49, 81 (1968). (9) J. C. Giddings, Sep. Sci., 8, 567 (1973). (10) L. F. Kesner, Electrical Field-Flow Fractionation, Ph.D. Thesis, University of Utah, 1974. (11) J. C. Giddings, Y. H. Yoon, K. D. Caldwell, M. N. Myers, and M. E. Hovingh, Sep. Sci., I O , 447 (1975). (12) C. Tanford, "Physical Chemistry of Macromolecules", John Wiley and Sons, New York, 1961, Chapters 6 and 7. (13) W.F. Harrington, P. Johnson, and R. H. Ottewill, Biochem. J., 62, 569 (1956). (14) "Handbook of Biochemistry", H. A. Sober, Ed., The Chemical Rubber Co., Cleveland, Ohio, 1968, p c-39. (15) E. 0. Fieid and J. P. P. O'Brien, Biochem. J., 60, 656 (1955). (16) J. R. Coivin. Can. J. Chem.. 30. 841 (1952). i17j R. E. Stauffer, in "Physical Meihods of Organic Chemistry", 26 ed., Part 1, A. Weissberger, Ed., Interscience, New York, 1956, p 65. (18) H. Svensson, Adv. frotern Chem., 4, 264-268 (1948). (19) T. Teorell, J. Gen. Physiol., 19, 917 (1936).
RECEIVEDfor review April 2,1976. Accepted July 23,1976. This investigation was supported by National Institutes of Health Grant GM 10851-19. LFK was supported by NIH predoctoral fellowship 5 FoL GM 41865.
High Speed Ion Exchange Chromatography of Proteins Shung-Ho Chang, Rodney Noel, and Fred E. Regnier" Department of Biochemistry, Purdue University, West Lafayette, Ind. 47907
Polymerization ~ t triglycidylglycerol f on the surface of controlled pore glass supports with a glycerylpropylsilylboarded phase produced a chromatographicallystable glycerol polymer to which stationary phase groups were attached. Dlethylaminoethyl (DEAE), methyldiethylaminoethyl (QAE), carboxymethyl (CM), and sulfonylpropyl (SP) ion exchange-supports of thls type with pore diameters exceeding 100 d were suitable for the separation of proteins. These pellicular supports were capable of withstanding pressures in excess of 5000 psi and linear flow rates to 5 mm/s. Separationspeeds 10 times faster than those of classical gel type supports were obtained when using comparable column length, particle slre, and elution protocol. Hemoglobin ion exchange capacities vary from approximately 40 mg/cm3 on 250-8 pore diameter supports to 20 mg/crn3 on 550-d pore material.
Proteins have been fractionated classically by chromatography on carbohydrate gel columns. These separations were achieved principally by steric exclusion, ion exchange, and
more recently by affinity chromatography. The success of carbohydrate gel supports in the separation of biopolymers is due primarily to: 1)their ability to imbibe large quantities of water and swell into a hydrophilic matrix, 2) their porosity to macromolecules, and 3) the ability of the hydrophilic carbohydrate matrix to stabilize sensitive biological compounds such as proteins. Unfortunately, the carbohydrate gel beds are sensitive to changes in pH, ionic strength, and pressure. This seriously limits the speed with which a resolution may be achieved and restricts the elution protocol. Elution times of 4-24 h with gentle gradients are common for the resolution of protein mixtures ( I ) . An additional disadvantage of many of the carbohydrate ion exchange supports is that they must be removed from the column for recycling and be repacked between separations. The preparation by Haller (2)of controlled porosity glass (CPG) that was porous to biopolymers and insensitive to changes in pH, ionic strength, and pressure provided a support matrix that appeared to have promise in the high speed resolution of proteins. Unfortunately, CPG irreversibly adsorbs and/or denatures many proteins and was found to be of lim-
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
1839
ited general utility in the resolution of proteins (3). It has recently been shown ( 4 ) that a chemically bonded surface layer of glycerylpropylsilane on the surface of glass or silica will overcome these problems of the adsorption and denaturation of proteins. This paper describes the preparation and use of totally porous pellicular ion exchange chromatography supports for proteins. These supports consist of a thin layer of skin of glycerol polymer bonded to the surface of CPG with ion exchange groups coupled to the hydrophilic polymer skin.
EXPERIMENTAL Apparatus. Chromatographic separations were carried out with a chromatographic system consisting of an Isco model 384 pumping system (Instrument Specialties Company, Lincoln, Neb.), a Disc model 706 sample injection valve (Disc Instruments Inc., Costa Mesa, Calif.), and a Perkin-Elmer model LC 55 detector. The columns were 4.8 mm i.d. X 0.25 in. 0.d. stainless steel with 10-p pore stainless steel inlet and outlet column terminators. All connecting tubing was 0.01 in. i.d. X 0.063 in. 0.d. stainless steel. Reagents. Glycidoxypropyltrimethoxy silane, triglycidoxyglycerol, diglycidoxyglycerol, allyglycidyl ether, and butadiene diepoxide were obtained from Polyscience, Inc., Warrington, Pa. Controlled porosity glass (CPG) was obtained from Corning Glass Works, Medfield, Mass. Boron trifluoride etherate and p-nitrophenyl acetate were purchased from Aldrich Chemical Co., Milwaukee, Wis. Soybean trypsin inhibitor, trypsin, chymotrypsin, chymotrypsinogen, hemoglobin, and myoglobin were obtained from Sigma Chemical Co., St. Louis, Mo. Procedures. Preparation of Glycerolpropylsilyl CPGISupports. The "glycerol" bonded supports were prepared by reacting CPG with a 10%aqueous solution of glycidoxypropyl trimethoxy silane at 95 "C for 2 h as described previously ( 4 ) . Oxirane Monomer Coating of Supports, Oxirane monomers were coated on supports by either a filtration or an evaporation process. In the filtration method, 50 ml of a 5% solution of oxirane monomer(s) in acetone was added to 10 g of support. After filtration, acetone remaining in the support pores was removed at room temperature in a fluidized bed drying apparatus. Monomer coating by evaporation was achieved by adding 20 ml of acetone containing 1 g of oxirane monomer(s) to 10 g of glycerylpropylsilyl CPG supports in an evaporating dish. After placing in a fume hood, the slurry was stirred intermittently with a spatula as the acetone evaporated. Removal of final traces of acetone was achieved in a fluidized bed. Fluidized Bed Polymerization. Ten grams of 37-74 p support coated with oxirane monomer(s) as outlined above were added to a 2-in. X 36-in. glass chromatography column fitted with a coarse glass frit. A nitrogen gas cylinder was connected to a 24/40 standard taper joint on the bottom of the column and the support bed was fluidized with a reverse flow of nitrogen. After all traces of organic solvents were removed, the nitrogen flow was temporarily interrupted and a 250-ml double necked round bottom flask containing 25 ml of BFa-etherate was fitted on the bottom of the column. Connecting the nitrogen stream to the second joint of this flask again fluidized the bed in addition to sweeping BFS-etherate vapor into the fluidized bed. Polymerization of oxirane monomers was completed in 15 min at 25 "C. Preparation of the D E A E Support. Ten grams of 37-74 p (250 A) glycerolpropylsilyl bonded CPG prepared according to the procedure outlined above was added to a 250-ml Erlenmeyer flask and treated for 18 h a t 90 "C with 100 ml of a solution containing 20% triglycidylglycerol, 40% diethylaminoethanol, and 40% dimethylformamide (v/v/v). Diethylaminoethanol serves as both a catalyst and reactant during polymerization. After filtration, the support was washed sequentially with 500 ml of water and 100 ml of acetone and then dried in vacuo. The support was then coated with 50 ml of a 5% solution of triglycidylglycerol in acetone by the filtration procedure. Polymerization was achieved in a fluidized bed as described above. Preparation of QAE Support. Five grams of DEAE support was added slowly to a 100-ml flask containing 50 ml of a 10%solution of CH3I in methanol. The slurry was heated a t 40 "C with occasional mixing for 15 h. After filtration, the support was washed consecutively with 100 ml of the methanol and acetone followed by drying in vacuo. Preparation of CM Support. Fifty ml of diethyl ether containing 250 mg of triglycidylglycerol and 250 mg of allylglycidyl ether were added to 10 g of 37-74 p (500-A pore diameter) glycerylpropylsilyl CPG. After coating the monomer(s) by the evaporation method, final removal of ether was achieved in the fluidized bed. Copolymerization 1840
0
of oxirane monomers was achieved in the fluidized bed with BF? etherate catalysis. Residual catalyst was purged with nitrogen and the support was removed from the fluidizer and washed with 500 ml of water. Conversion of allyl residues in the organic skin to a carboxymethyl (CM) stationary phase support was achieved by oxidation with 200 ml of 8 mM sodium metaperiodate, 2 mM potassium carbonate, and 0.134 mM potassium permanganate ( 5 ) .Removal of oxidants by filtration was followed by consecutive 500-ml washes with 1M sodium bisulfite and water. A final wash with acetone and vacuum drying completed the preparation of the support. Preparation of Sulfonyl Propyl ( S P )Support. Ten grams of 37-74 p (500 A) triglycidylglycerol-aliylglycidyl copolymerized support from the CM preparation were added to 200 ml of 1 M sodium bisulfite. Oxygen was slowly bubbled through the slurry for 4 h at 25 "C yielding the sulfonic acid derivative (6). After filtration, the support was washed with water and the acetone and air dried. Ion Exchange Capacity Measurements. The method for determining hemoglobin ion exchange capacity has been described previously (7). Preparation of Columns.All columns were packed with dry support by the "tap-filled" method (7). Preparation of Creatine Phosphokinase ( C P K ) Isoenzymes. A mixture containing the three CPK isoenzymes was prepared by mixing two partially purified CPK preparations. The first CPK preparation, yielding only CPK3 activity, was isolated from beef heart according to the procedure of Henry et al. (8).The second preparation, containing approximately equal portions of CPKl and CPK2, was isolated from beef brain by a procedure similar to that of Kuby et al. (9). Protein and Enzyme Recovery. Recovery of creatine phosphokinase, lactic dehydrogenase, alkaline phosphatase, and trypsin enzyme activities from chromatography columns were determined by placing 270 pl of enzyme solution in the initial buffer on a 0.48 X 50 cm DEAE glycophase1CPG column and eluting with a 20-min gradient to 100% of the final buffer a t a flow rate of 3 ml/min. Buffer concentrations and pH for each enzyme are in Table 111. The total eluent was collected and enough final buffer added to make a final volume of 70 ml. Controls were treated in the same manner except that the DEAE column was removed from the apparatus. Creatine phosphokinase was assayed according to the Rosalki method (IO)and the three remaining enzymes by procedures outlined in the Worthington Enzyme Manual (11). Hemoglobin recovery from DEAE supports was determined in a batch assay. One hundred mg of DEAE support was initially washed with 3 ml of 0.01 M Tris.hydrochloride buffer (pH 8.8) and followed by treatment with 30 mg of hemoglobin in 3 ml of the above washing buffer. The hemoglobin solution and DEAE support were vortex mixed five times in a 30-min interval a t room temperature. Centrifugation, decantation, and reequilibration with several 3-ml portions of the 0.01 M Tris-hydrochloride buffer (pH 8.8) removed unbound hemoglobin. Hemoglobin was released from the DEAE support with 0.4 M Tris-hydrochloride buffer (pH 3.5). After three washings with 3-ml portions of the pH 3.5 buffer, the pooled supernatant solutions were diluted to a volume of 10 ml and the hemoglobin recovery was determined spectrophotometrically a t 410 nm.
THEORETICAL The bonding of stationary phases to inorganic supports was accomplished by forming a thin layer of glycerol polymer on their surface that was attached to the support through an organosilane coupling agent with the stationary phase bound to the inert polymer matrix. This composite has the general chemical formula =SiCH2CH2CH2R1PS where R1 is a glycerol polymer and P, is the stationary phase. The primary function of the polymer is to provide an organic skin that is hydrophilic, neutral, stable, and will not partition proteins by itself. Synthesis of pellicular supports was achieved by first coating the surface of glycerolpropylsilylbonded CPG (4)with a thin layer of oxirane monomer followed by polymerization. By increasing the amount of oxirane monomer(s) with multiple oxirane functional groups, the degree of cross-linking in the polymer could be increased. Since many proteins are absorbed on hydrophobic materials, only hydrophilic oxirane monomers were used in polymer coatings. A series of oxirane monomers used in polymer formation are shown in Table I. During the course of polymerization, the hydroxyl groups of the glycerolpropylsilyl-CPG (4) become incorporated into
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
Table I. Oxirane Monomers Used in Support Preparation Oxiranes/molecule
Monomer
FHz-
0 / \
OCHzCHCHz
I
3
Trigly cidyl glycerol
Post polymerization function
Structure
0
Hydrophilic support layer
r-OCHzC / \F
CHz--OCH,CHCH2 0
// CHzOCHzCHCHz
2
Dig1ycidyla glycerol
Hydrophilic support layer
CHOH
I
/\
CHzOCHZCHCHz 0 /\ CH2-OCH,CHCH, 0 /\ CHz-OCHzCHCH,
Hydrophilic support layer
Diglycidyl ethylene glycol
2
Butadiene diepoxide Allyl glycidyl ether
2
0 0 /\ /\ CH&HCHCH,
Hydrophilic support layer
1
/0\ CH2=CHCHz0CH2CHCHz
Intermediate in preparation of cation exchanger
UThis compound is probably a mixture of
I
cy,&'-
and a$-diglyceryl ethers with the former predominating.
the polymer matrix and couple the polymer to the support as shown in reaction 1.
oxirane containing stationary phase or stationary phase derivative has the general chemical formula
lo\
OH
CHz- CHCH,OP,
I
sSiCH,CH,CH,0CH2CHCH20H
and is incorporated into the glycerol polymer matrix as shown in reaction 2.
O /\ /"\ + (CHZ-CHCH@CH2)2CHOCH,CHCH,
BF, -t
OH I I
OH
I
?'I
=SiCH2CH,CH,0CH2CHCH,0H
~SiCH,CH,CH,-f-OCH,CHCH, j,+OCH,CH-CH,-f;; (1)
/O\ + (CH2-
O /\
CHCH,OCH&CHOCH,CH-CH,
O/
Although triglycidyl glycerol was used in the example, any of the multifunctional oxirane monomers listed in Table I may be used. I t will be shown later that catalysts other than BF3 may also be used. The most critical part of the process is to establish a thin, uniform surface layer of oxirane monomer that will not fill the pores of porous supports. This is accomplished by either slurry (12) or filtration (13) coating of the oxirane monomer from a volatile organic solvent followed by fluidized bed drying. As the volatile solvent evaporates in the fluidized drying apparatus, a film of oxirane monomer is deposited on the surface of the support. Polymerization with BF3 is achieved by sweeping the volatile polymerization catalyst into the fluidized drying apparatus as the coated packing material is being tumbled in a stream of nitrogen. Tumbling the support during polymerization with a gaseous catalyst leaves the surface film of oxirane intact while preventing the aggregation of support particles. Stationary phase (P,) groups may be introduced into the polymer matrix in two ways: 1)copolymerization and 2) coupling during polymerization. In the copolymerization method, an oxirane monomer containing the stationary phase (P,) or a functional group that will subsequently be converted to a P, and one of the multifunctional oxirane monomers in Table I are coated on the support together and copolymerized. The
I
O /\ + CH,-CHCHjOP. OH
I 3~ S i C H , C H , C H , - f - O C H 2 C H C H 2 ~ O C H , C H C H C H ~ ~ P s ) o (2)
Copolymerization is carried out in a fluidized bed with BF3 catalysis in all cases. Stationary phases (P,) that contain a nucleophilic functional group may also be added to oxirane monomers during the course of polymerization as indicated in reaction 3. The group X represents the nucleophile on the stationary phase before bonding. The addition OH
I
~SiCH,CH,CH,0CH2CHCH20H
?\ ?\ + + (CH&HCH,OCH&CHOCH,CHCH,
-
X-P,
OH
/"\
I
SiCH2CH2CH2f-OCH2CHCH,-f;;;tOCHCHCH,~XP,), 13)
of a nucleophile to the oxirane in this case essentially terminates its participation in further cross-linking reactions. Catalysts necessary for this type of reaction vary with P, and are discussed more specifically in the Experimental section.
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
1841
Table 11. Ion Exchange Capacities and Microanalysis of Glycophase Supports Inorganic support
Particle size, p
Pore diameter, A
Surface area, m2/g
Stationary phaseb
Organic carbon, % by w t
Ion exchange capacity mg/cm3c
170 150 CM 6.69 31 CPS 20-30 250 130 CM 5.71 48 CPG 37-74 550 70 CM 4.09 27 CPG 37-74 CM 4.21 25 CPS 37-74 500 ... 550 70 SP 4.03 18 CPG 37-74 CPG 74-128 100 170 DEAE 6.95 34 250 130 DEAE 5.97 45 CPG 74-128 DEAE 4.36 20 CPG 74-128 550 70 6.37 28 CPG 74-128 100 170 QAE 4.26 36 CPG 74-128 250 130 QAE aCPS and CPG designate controlled porosity silica and glass, respectively. b The abbreviations CM, SP, DEAE, and QAE represent carboxymethyl, sulfonylpropyl, diethylaminoethyl and quaternary anion exchange group, respectively. C The ion exchange capacity is expressed in mg hemoglobin bound per cm3 of support. Specific details for determining hemoglobin ion exchange capacities are found in the Experimental section.
Table 111. Protein and Enzyme Recovery from DEAE Glycophase Supports Protein
Initial buffer
Final buffer
Recovery, %
Hemoglobin Creatine phosphokinase
0.01 M Tris, pH 8.8 0.05 M Tris, 1 mM mercaptoethanol,
95a 96b
Lactic acid dehydrogenase Alkaline phosphatase Trypsin
pH 8.0 0.025 M Tris, pH 8.0 0.05 M Tris, pH 8.0 0.05 M Tris,
0.4 M Tris, pH 8.8 0.05 M Tris, 0.3 M NaC1, 1 mM mercaptoethanol, pH 8.0 0.05 M Tris, 0.2 M NaC1, pH 8.0 0.05 M Tris, 0.3 M NaCl, pH 8.0 0.05 M Tris, 0.3 M NaC1, pH 8.0
pH 8.0 a
9 9b 91b
94 b
Recovery of total protein. b Recovery of enzyme activity.
In the case of the Preparation of DEAE Support, the hydroxyl of diethylethanol amine serves as the nucleophile while the tertiary amine is the catalyst. Since the immobilized polymer layer is composed of the simple sugar alcohol glycerol, we have chosen the name “Glycophase” to designate the bonded phase. Specific supports, such as the diethylaminoethyl ion exchange support, will be referred to as DEAE glycophase/CPG.
RESULTS Our studies indicate that when a thin layer of glycerol polymer containing ion exchange groups is bonded to controlled porosity glass or silica according to the procedures outlined above, a family of ion exchange supports is produced that is suitable for the high speed chromatography of proteins. These supports have ion exchange capacities approaching those of classical carbohydrate-gel supports, equilibrated rapidly, and show good resolution of macromolecules at mobile phase velocities of 1mm/s. The hemoglobin exchange capacities for a series of supports is shown in Table 11. Ion exchange capacities are reported in mg protein/cm3 of support so that these results may be compared with data on commercially available carbohydrate-gel type ion exchange supports. It will be noted that ion exchange capacity varies with the pore diameters of supports. Since ion exchange groups are bonded to the support surface and less than 1%of the surface area of a totally porous inorganic support is on the outside of a support particle, it is necessary for a protein to penetrate the support pores to be ion exchanged. Penetration of the 100-A pore diameter support by the large hemoglobin molecule is incomplete and therefore the macromolecule is not able to reach all of the ion exchange groups. The 250-A support on the other hand is totally penetrated by hemoglobin and has a higher ion exchange capacity even 1842
though its surface area is less than that of the 100-A pore diameter support. The effect of a decrease in surface area on ion exchange capacity of a material that is totally penetrated by solute is seen in the 550-A support. It is obvious that ion exchange chromatography of macromolecules on supports that have pore diameters similar to the molecular dimensions of solutes is a combination of ion exchange and steric exclusion chromatography. The nature of the inorganic support has little influence on ion exchange capacity if the pore diameter and surface area are the same. The hemoglobin ion exchange capacities of both controlled pore glass and silica with 500-A pores and a CM coating were consistently 26-30 mg/cm3. Protein recoveries from these ion exchange supports in the several cases tested were good. It is noted in Table I11that the recovery of protein and enzyme activity from the DEAE columns exceeded 90% in all cases. This indicates the polymer coating eliminates most of the irreversible absorption and protein denaturation problems found on native CPG supports. Degradation of column efficiency with increasing mobile phase velocity for the DEAE ion exchange support is seen in Figure 1.This plate height ( H )vs. mobile phase velocity (y) curve is very similar to the one obtained on glycerolpropylsilyl/CPG supports used for the high speed gel filtration of proteins ( 4 ) . It has been our experience that column efficiencies for proteins are lower than those for small molecules. Although there are no data to explain this phenomenon, it is probably due to the fact that proteins have 10- to 100-fold smaller diffusion coefficients than small molecules. Band spreading will be enhanced by the much slower rate of diffusion of proteins in stagnant mobile phase pools within support pores and the slower rate of mass transfer from the mobile phase to support particles. Since the rate of solutr transfer
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
DEAE
- GlycophoseICPG
I"
.' 74 - 128p,250
E,
SAMPLE:
Commercial Soybean Trypin Inhibitor
PACKING:
CM Glycophooe/CPG 2501 pore. 3 7 - 7 4 ~
COLUMN:
4 . 8 x 4 0 0 mm SS
SOLVENT:
s
A =0.005M Acelale. pH 4.0
B = 0 . 0 0 5 M Acetate, 0.6 M NaCl. pti 4 . 0
N
a
I
FLOW RATE! 5 m m /sec.
W v)
z
2
PRESSURE:
75 prlq
DETECTOR:
UV 2 5 4 nm. 0.02 AUFS
v)
W
a a
0 I-
O W I-
W
/
P
TIME (min.) Figure 2. Chromatography of commercial soybean trypsin inhibitor on 37-74 w, 250-A pore, CM GlycophaseICPG
I
2
3
4
The sample was gradient eluted from a 4.8 mm X 400 mm stainless steel column at a linear flow rate of 5 mm/s and a pressure of 75 psi. Solvent A was 5 mM sodium acetate buffer (pH 4.0) while solvent B was 5 mM sodium acetate buffer (pH 4.0) containing 0.6 M sodium chloride. The peak eluting between 15 and 25 min contained all soybean trypsin inhibitor activity
5
u ( m m /set.) Figure 1. Effect of mobile phase velocity on efficiency of DEAE glycophase supports
loot
These values are based on the isocratic elution of myoglobin from a 4.8 mm X 600 cm column with 0.05 M Tris-HCI (pH 8.0) containing 0.01 M sodium chloride
between mobile and stationary phases would be expected to be much slower for macromolecules, decreased mobile phase velocities would be required to allow equilibration of solutes. By using the equation for reduced mobile phase velocity (14) v = yd,/D, and diffusion coefficients of loM5and for small solutes and proteins, respectively, it may be calculated that a comparable mobile phase velocity for the protein columns should be 10%that used for small molecules; where y is mobile phase velocity, d, is particle diameter, and D, is the solute diffusion coefficient. This is generally what we have observed. After an ion exchange gradient elution, columns were recycled by one of two procedures. When there was a large difference in the pH and ionic strength of the initial and final buffers, columns were recycled by retracing the elution gradient and pumping several column volumes of the initial buffer through the column. The second recycling technique consisted of switching at the end of an analysis directly back to the initial buffer. When the pH did not vary more than 2 units, recycling was complete with the direct switchback procedure after the passage of 5 column volumes of the initial buffer. While operating between the limits of pH 4 and 8, one 250-A pore diameter DEAE glycophase/CPG column was operated through 100 cycles without loss of resolution. Elution of columns with pH 8,0.5 M sodium phosphate buffers at 100 OC for 48 h without sample introduction caused a decrease in column bed volume. Carbon-hydrogen analysis indicated that the relative percentage of organic polymer on the support increased during this settling process. This is interpreted to mean that the inorganic surface is eroding under the organic polymer layer. If this is true, the ultimate stability of these inorganic-organic polymer matrices is dependent on the stability of the inorganic support material in a particular mobile phase. Operation of these ion exchange supports in acidic media to pH 2 posed no apparent stability problem.
II
H)
0 I
1
I I
I
I
I
I I
I
E U
l 0
m cu
I
W v)
I
I I 0
I I
v)
0 I I 0
W
U
m Iz
I
z
B
? ?
I
W
3
0
v)
I-
z W 0
U
w
a
1
TIME (min) Figure 3. Separation of a commercial chymotrypsin preparation on 74-128 w , 250-A pore, SP GlycophaselCPG A 4 mm X 300 mm stainless steel column was gradient eluted at a linear velocity of 1 mm/s with 50 psi of column inlet pressure. Solvent A was 10 mM sodium phosphate buffer (pH 8.0) while solvent B was 10 mM sodium phosphate buffer (pH 8.0) with 0.5 M sodium chloride. The peak eluting between 12 and 15 min contained all chymotrypsin activity
The elution profile for a commercial sample of soybean trypsin inhibitor on the 250-A pore diameter CM Glycophase/CPG support is seen in Figure 2. The compound eluting from the column in 20 min was shown to be the inhibitor by its trypsin inhibitor activity. This elution order is the same as that obtained from a soybean acetone powder on CM cellulose in 10 h ( 2 5 ) . Further analyses with this column of amylase, immunoglobulins, myoglobin, and hemoglobin samples were accomplished. Fractionation of myoglobin and hemoglobin preparations according to elution protocols for
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
1843
Ioc
I
SAMPLE:
Crude Trypsin
PACKING:
DEAE GlycophaseICPG 250 & Pore, 37-74p
COLUMN:
4 . 8 x 500 mrn SS
SOLVENT:
A :0.05 M Tris, pH :8.0 8 = 0.05 M Tris, 0.3 M NaCl,
I00 psi
DETECTOR:
254 nm
Protein Mixture PAE GlycophaseICPG 250 A Pore, 74 l28p
COLUMN:
4 . 8 x 600 mrn
SOLVENT:
A = 0.02 M Tris. pH 8 . 0 B = 0.05 M Tris. 0.3 M NoCI, pH = 7.5
pH = 8.0
FLOW RATE] 4 . 5 rnmIsec
PRESSURE:
SAMPLE: PACKING:
-
SS
FLOW RATE: 4.5 mm/sec
PRESSURE:
100 psi
DETECTOR:
UV 254 nm
PEAK IDENTITY:
: Chymotrypsinogen b = Myoglobin c = Myoglobin + Hemoglobin d = Albumin
o
TIME (rnin.) Figure 6. Resolution of proteins on a QAE Glycophase column
IO
20
TIME (min.) Figure 4.
Separation of the components in a crude commercial trypsin
The 4.8 mm X 600 mm stainless steel column packed with 74-128 p, 250-A pore, QAE Glycophase/CPG was gradient eluted at a linear velocity of 4.5 mm/s and a pressure of 100 psi. Solvent A was 20 mM Tris buffer (pH 8.0) while solvent B was 50 mM Tris (pH 7.5) with 0.3 M sodium chloride. Peak identity: a = Chymotrypslnogen, b = Myoglobin, c = Hemoglobin, d = Albumin
sample The 4.8 mm X 500 mm stainless steel column packed with 27-74 p, 250-A pore DEAE Glycophase/CPG was operated at a linear velocity of 4.5 mm/s and a pressure of 100 psi. Solvent A was a 50 mM Tris buffer (pH 8.0) while solvent B was 50 mM Tris buffer (pH 8.0) with 0.3 M sodium chloride. The peak indicated by the arrow contained all trypsin activity SAMPLE: Bovine CPK Isoenzymes PACKING: D E A E Glycophase/CPG COLUMN: 4.8 x 6 O O m m SS SOLVENT: A = 0.05M Tris ( p H 7.5), 0.05M NoCl, 0.001M Mercaptoethanoi 8 = 0.05M Tris (pH7.51, 0 . 3 M NoCI, 0.00iM Mercaptoethanol FLOW R A T E : 2 mm/sec. PRESSURE:
250 psi
TIME ( m i d Figure 5.
Separation of bovine creatine phosphokinase isozymes
The 4.8 mm X 600 mm stainless steel column packed with 37-74 p, 250-A pore, DEAE Glycophase/CPG was gradient eluted at a linear velocity of 2.2 mm/s and a prossure of 50 psi. Elution solvent A was a solution of 5 mM Tris buffer (pH 7.5), 50 mM sodium chloride, and 1 mM mercaptoethanol while solvent B was a solution of 50 mM Tris buffer (pH 7 4 , 0.3 M sodium chloride, and 1 mM mercaptoethanol.The elution order of the three isoenzymes Is MM, MB, and BB, respectively
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CM cellulose (16)was also achieved on the 250-A CM Glycophase/CPG support in 20 min. The chromatography of chymotrypsin on 250-A pore diameter SP Glycophase/CPG is shown in Figure 3. Enzymatic hydrolysis of p-nitrophenylacetate indicated that the final peak eluting from the column was chymotrypsin. This column was also used in the separation of chymotrypsinogen and trypsin from impurities in less than 20 min. Analysis of a commercial trypsin sample on 250-A pore diameter DEAE Glycophase/CPG is shown in Figure 4.Trypsin activity was found in the second peak (indicated by arrow). Further use of this column in the resolution of the three creatine phosphokinase isoenzymes from bovine tissue is shown in Figure 5. This mixture was prepared by recombining the partially purified individual isoenzymes. Enzymes were detected by a continuous enzyme activity monitoring system ( 17). The MM isoenzyme elutes first followed by the MB and BB isoenzymes, respectively. The identity of the individual isoenzyme was confirmed independently by collecting fractions from the ion exchange column and electrophoresing them on cellulose acetate strips vs. authentic standards. The electrophoresis of the isoenzymes was carried out by the procedure described by Henderson (18). The separation of a protein mixture containing commercial chymotrypsinogen, myoglobin, hemoglobin and albumin on 250-A pore diameter QAE Glycophase/CPG is shown in Figure 6. This strong anion exchanger could be particularly good for the separation of proteins with high PI values based on the fact that the separation of chymotrypsinogen and myoglobin is much more difficult on DEAE Glycophase/CPG than QAE.
CONCLUSIONS It may be concluded that through the use of forced flow and rigid supports, ion exchange chromatographic resolution of protein mixtures is achievable in 30 min or less. Controlled porosity glass particles provide a suitable matrix for the covalent attachment of ion exchange groups when the inorganic support has been coated with a hydrophilic polymer layer that deactivates the inorganic surface and sterically excludes biological macromolecules from contact with unreacted surface
ANALYTICAL CHEMISTRY, VOL. 48, NO. 13, NOVEMBER 1976
silanols. Rapid analysis of a variety of proteins of industrial, pharmaceutical, clinical, and general research interest is possible using commercial liquid chromatographic equipment.
LITERATURE CITED E. Heftmann, "Chromatography", Reinhold Publishing Corporation, New York, 1967, pp 405-428. W. Haller, "Material and Method for Performing Steric Separations", U.S. Patent 3,549,524. C. W. Hiatte, A. Shelokov, E. J. Rosenthal, and J. M. Galimore, J. Chromatogr. 56,362 (1971). F. E. Regnier and R. Noel, J. Chromatogr. Sci., in press. E. Von Rudloff, Can. J. Chem., 43, 2260 (1965). J. D. Roberts and M. C. Ccserio. "Basic Principles of Organic Chemistry", W. A. Benjamin, New York, 1964, p 762. S.H. Chang, K. M. Gooding and F. E. Regnier, submitted to J. Chromatogr. Sci. P. D. Henry, R. Roberts, and B. E. Sonel, Clin. Chem. ( Winston-Salem, N.C.), 21,884 (1975).
H. J. Kentel, K. Okabe, H. K. Jacobs, F. Ziter, L. Maland, and S. A. Kuby, Arch. Biochem., 150,648 (1972). S.B. Rosalki, J. Lab. Clin. Med., 69,696 (1967). "Worthington Enzyme Manual", Worthington Biochemical Corporation, 1972. Howard Purnell, "Gas Chromatography", Wiley, New York, 1962, p 240. E. C. Horning, E. A. Moscatelli, and C. C. Sweeley, Chem. lnd. (London), 751 (1959). J. C.Giddings, "Gynamics of Chromatography Part K", Marcel Dekker, New York, 1965, p 58. Y. Birk, A. Gertler, and S. Khalef, Blochem. J., 87,281 (1973). A. Skeson and H. Theorell, Arch. Biochem. Biophys., 91,319 (1960). S.H. Chang and F. E. Regnier, in preparation. D. A. Nealson and A. R . Henderson, Clin. Chem. ( Winston-Salem, N.C.), 21, 392 (1975).
RECEIVEDfor review April 12,1976. Accepted July 26,1976. Parts of this work were presented at the 26th and 27th Pittsburgh Conferences on Analytical Chemistry and Applied Spectroscopy.The work was supported in part by a grant from Corning Glass Works, Corning, N.Y.
Universal Detector for Liquid Chromatography Based upon Dielectric Constant Leon N. Klatt Analytical Chemistry Division, Oak Ridge National Laboratory, Oak Ridge, Tenn. 37830
A liquid chromatography detector based upon dielectric constant is described. The detector employs a modular phaselocked-loop circuit with its operating frequency determined by the dielectric constant of the column effluent. Readout is the shift in frequency resulting from changes in solution dlelectric constant as a solute band elutes from the column. Response is independent of intrinsic solvent conductivity and dielectric loses associated with dipole-dipole relaxation. The detector has excellent stability and reproducibility, negligible sensitivity tto variations in solvent flow rate, and the capability of operating without any restrictions on solvent dielectric constant. Separations employing low and high dielectric constant eluants were monitored. An average detection limit of 0.0005 A€ was measured in low dielectric constant media. Comparison with the uv and refractive index monitors indicates that the dielectric constant detector should find numerous applications in liquid chromatography.
Liquid chromatography detectors are classified either as universal or specific; the universal detector responds to a change in some bulk property of the mobile phase as the solute elutes from the column while the specific detector responds to some unique property of the solute. Since dielectric constant is an electrical property possessed by all materials, a liquid chromatography monitor based upon dielectric constant is classified as a universal detector. The first use of dielectric constant as the basis for detection in liquid chromatography was probably made by Troitskii ( I ) . The response consisted of a change in an audio tone upon location of an absorption zone on an alumina column. Monaghan and co-workers ( 2 ) described the use of a heterodyne chemical oscillometer for chromatographic detection. The capacitor plates were separated from the solution by the insulating glass walls of the detector cell. This same instrumental concept was used by Baumann and Blaedel ( 3 ) and
Johansson et al. ( 4 ) to monitor a change in solution conductivity as a solute band passed a detector head attached to the chromatographic column. Based upon the equivalent circuit for these systems (5),one senses a combination of capacitance and conductance with the relative magnitude of each dependent upon the operating frequency of the oscillometer and the conductivity of the media. Vespalec and Hana (6) reported a capacitance detector for liquid chromatography that used a detector cell with direct contact between the capacitor plates and the column effluent. The instrument operated at 18 MHz, and the cell formed the capacitor in the tuned network that determined the oscillator operating frequency. Readout was obtained via a heterodyne technique. The detector cell had to be altered to permit operation with mobile phases of different dielectric constants (7). Use of the detector cell in the resonant network of the oscillator precludes operation with high dielectric constant solvents because the energy losses to the solution from dielectric relaxation quench the oscillations. Erbelding (8)described a very simple circuit based upon a triangular wave generator whose output was the input to a difference-differentiator containing a reference capacitor and the capacitance detector cell. Under ideal conditions, the amplitude of the square wave output from the differencedifferentiator is proportional to the capacitance difference between the detector cell and the reference capacitor. This circuit also was unable to operate with high dielectric constant solvents, although sufficient energy to compensate for dielectric losses was available. Because solution conductivity and dipole relaxation appear as a resistive component in parallel with the capacitance, the transfer function of the difference-differentiatoris dependent upon solvent dielectric constant producing an output waveform that is not uniquely related to capacitance and results in the detector's failure to operate with high dielectric constant solvents. The instrument described herein overcomes many of the difficulties inherent in previously described dielectric constant detectors and possesses sufficient sensitivity for use in modern liquid chromatography.
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