Anal. Chem. 1987, 5 9 , 1021-1027
1021
High-Performance Capillary Electrophoretic Separation of Bases, Nucleosides, and Oligonucleotides: Retention Manipulation via Micellar Solutions and Metal Additives A. S. Cohen,' S. Terabe? John A. Smith,3and B. L. Karger*'
Barnett Institute and Department of Chemistry, Northeastern University, Boston, Massachusetts 02115, Department of Industrial Chemistry, Kyoto University, Kyoto, Japan, and Department of Molecular Biology and Pathology, Massachusetts General Hospital and Department of Pathology, Haruard Medical School, Boston, Massachusetts 02114
High-performance Capillary electrophoretlc separatlon of bases, nucleosldes, and nucleotides has been achleved wlth the use of sodium dodecyl sulfate, SDS, mlcelles. Rapld Separations have been demonstrated wlth extremely narrow peak widths. Slnce the bases and nucleosldes are uncharged at the pH of operation (pH 7), separation Is a result of dlfferentlal partltlon wlthln the Interior of the mlcelle; the more hydrophobic the species, the larger the partltlon coefficient and the larger the retention. Ollgonucleotldes are negatlvely charged and can be separated wlthout SDS mlcelles; however, the tlme window is narrow and separation of complex mixtures llmlted. The combination of low concentrations of divalent metals and SDS mlcelles leads to a slgnlflcant enhancement of the tlme window and good separation of 011gonucleotldes with high theoretlcal plate counts. The metal Ion is electrostatically attracted to the surface of the micelle and dlfferentlal metal complexation of the ollgonucleotldes wlth the surface of the mlcelle leads to separatlon of complex mlxtures. As an example, 14 out of a mlxture of 18 oligonucleotides of 8 bases, each with a different sequence, is achleved in less than 30 mln by uslng a buffer system containing zinc and SDS mlcelles.
The use of capillaries for the high-performance electrophoretic separation of complex mixtures is an area currently under rapid development (1-6). The methodology often yields extremely sharp peaks, with bandwidths equivalent to 105-106 theoretical plates or greater. Since the fused silica capillaries employed have a negative zeta potential at their walls, unless suppressed bulk osmotic flow is toward the negatively charged electrode (8). Even negatively charged solutes move in this direction, however, at a slower velocity than the bulk flow itself. The high efficiency is, in part, a consequence of the uniform osmotic flow profile across the capillary (9) as well as the high potential (assuming axial diffusion controls and band broadening) (10). The purpose of this paper is to explore the capillary electrophoretic separation of nucleic acid constituents-bases, nucleosides, and oligonucleotides. The use of this approach to nucleic acid chemistry is potentially beneficial, given the high resolving power and low sample size requirements. Chromatographic (e.g., ion exchange (11)and reversed phase (12))and open bed electrophoretic procedures (13) are widely used today to separate nucleic acid constituents, especially oligonucleotides. There is one report on the capillary electrophoretic separation of nucleotides (14). A second purpose of this work is to increase the selectivity control of the capillary electrophoretic method through the
(a,
Northeastern University.
Kyoto University.
Harvard Medical School.
use of anionic micelles from sodium dodecyl sulfate, SDS. One of us has already shown that uncharged solutes can migrate differentially in the capillary via partition of the solute into the interior of the micelle (15). Sharp peaks are obtained by this approach. Since the bases and nucleosides are also uncharged at the pH of the buffer, we have taken advantage of differential micelle partition to achieve separation. Oligonucleotides are negatively charged, and some separation on the basis of electrophoretic mobility differences is possible. (Note also that these species will not adsorb to the negatively charged fused silica capillary walls.) However, we have found enhanced selectivity of these solutes via differential complexation with metals added to the buffer system containing SDS micelles. The divalent metals examined-Cu(II), Zn(II), and Mg(I1)-are known to be electrostatically attracted to the surface of negatively charged micelles (16). Oligonucleotide complexation with metals associated with the micelle affects migration and thus separation. By this approach, selectivity can be conveniently manipulated in the high-performance capillary electrophoretic separation of oligonucleotides. EXPERIMENTAL SECTION Apparatus. Electrophoretic separation of bases, nucleosides, and oligonucleotides was performed in a fused silica tubing 0.05 mm i.d. (Scientific Glass Engineering, Ringwood, Victoria, Australia) with various column lengths from 500 to 850 mm, depending on the experiment. A regulated high-voltage dc power supply, Model LG-40R-35for bases and nucleosides, or Model LG-30R-5 for oligonucleotides (Glassman, Whitehouse Station, NJ) which could deliver high voltage up to 30 kV was used to produce the potential across the capillary. UV detection was employed for bases and nucleosides (Jasco Uvidec 100 11, Tokyo, Japan) and for oligonucleotides (Soma S-3702, IR&D, Kingston, MA). The detection wavelength was 260 nm throughout. Both detectors were modified as described elsewhere (4). The tubing and detector were placed in a thermostated air bath at 35 "C for the base, and the nucleoside separations and the tubing was thermostated at 25 "C using a liquid cooling system (Lexacal Model Ex-100DD with FTC-350A, Neslab Instruments, Inc., Portsmouth, NH) for the oligonucleotide separations. A Nelson Analytical Model 762SB A/D interface (Cupertino, CA) attached to an IBM PC/XT was used to record the results and to process data for the oligonucleotides. Materials. Bases and nucleosides were purchased from Nakari Chemicals (Kyoto, Japan) or Kojin (Tokyo, Japan) and used without further purification. Sodium dodecyl sulfate (SDS) was of protein research grade (Nakarai Chemicals, Kyoto, Japan, or Schwarz/Mann Biotech, Cambridge, MA). The lower molecular weight polythymidines (up to six bases) were purchased from Sigma (St. Louis, MO); higher molecular weight oligonucleotides were synthesized with an Applied Biosystems 380A DNA synthesizer (Foster City, CA). The water was deionized and triply distilled. The other reagents were A.R. purity grade. All buffer solutions were filtered through a Nylon 66 filter unit of 0.2-pm pore size (Schleicher and Schuell, Keene, NH). The oligonucleotide samples were kept frozen at -20 "C, and working sample solutions were stored at 4 "C.
0003-2700/87/0359-1021$01.50/0 0 1987 American Chemical Society
1022
ANALYTICAL CHEMISTRY, VOL. 59, NO. 7, APRIL 1, 1987
Procedure. Capillary tubes were filled with the desired buffer using a 100-pL gas-tight syringe (Hamilton Co., Reno, NV). Both ends of the tube were then dipped into separate 5-mL reservoirs filled with the same buffer. The end in which samples were introduced was connected with platinum electrodesto the positive high voltage. The reservoir at the detector end was connected with platinum electrodes to ground. Samples were introduced either by electroinjection or by siphoning at a concentration level of 0.5-1.0 mg/mL. The migration of an uncharged species towas determined from the solvent peak or injection of a trace of methanol. The migration of the micelle t,, was determined by addition of the dye Sudan I11 to a micelle solution. The dye was assumed to be fully partitioned within the micelle, so that measurement of the migration time of the dye corresponded to tmc.
Before each run, the capillaries were purged with 100 pL of 0.1 M NaOH followed by 250 pL of triply distilled water. Care was taken to equilibrate the capillary with buffer prior to operation. The reproducibilityof retention was better than 1%relative standard deviation from run to run and better than 3% from day to day. Moreover, it was found that reproducibility remained at 3% from capillary tube to capillary tube within the same batch of fused silica capillaries. RESULTS AND DISCUSSION Solute Migration. Electroosmosis in a capillary is the bulk on the flow resulting from the charge or zeta potential, walls. The electroosmotic velocity, ueo,can be obtained from the migration time of a neutral species or solvent peak t o and can be expressed as (17)
cl,
where t is the dielectric constant, is the viscosity of the liquid medium, E is the electric field strength, and 1 is the length of capillary to the detector cell. Since E = VfL, where V is the voltage drop across the capillary of length L (note: 1 and L are not the same), eq 1 can be recast as ueo
= /*eo*V/L
(2)
where peo* is the electroosmotic mobility of the fluid. (pea* is used rather than peo in order to emphasize that the electroosmotic mobility is not a constant but is a function of the composition of the fluid and the zeta potential of the walls.) Since E is proportional to I , the current due to transport of charge by the fluid (15),we can rewrite eq 1 as
(3) where r is the radius of the capillary and x is the bulk conductance of the solution. Equations 1-3 indicate the various parameters that can affect urn, one of the most important being the zeta potential on the capillary walls. Analogously to eq 2, the electrophoretic velocity, uep, can be written as uep
=
/*epV/L
(4)
where pep is the electrophoretic mobility of a given species. The velocity of a charged solute is then simply 0,
=
ueo
+ uep
=
bee* + Mep)V/L
(5)
Note that if the solute migrates electrophoretically in a direction opposite to the bulk flow (as would be the case for negatively charged species with negatively charged fused silica capillaries), then u, would be smaller than u,, (Le., ueP would be negative). The migration time of the solute t, can be written from eq 5 as
An analogous expression to eq 6 can be written for the mi-
gration time of a micelle, t,,, such as with SDS. When neutral solute partitions within a micelle, the migration time of that species can be expressed as ( 4 ) ts
= (1
+ (to/tmc)R’) t o + & I
(7)
where 6’ is the capacity factor for partition of the solute between the micelle and the aqueous phase
where nmc/neqis the mole ratio of solute in the micellar and aqueous phases, respectively, K is the distribution coefficient, and V, and Vaqare the volumes of the micellar and aqueous phases, respectively. The range over which the solute can migrate is between t oand t,,, and this range, along with the efficiency of the process, determines the peak capacity. Analogously to chromatography, resolution can be expressed as (15)
where N is the number of theoretical plates and a is the separation factor 6 f 12; where subscripts 1 and 2 refer to the first and second components, respectively. Optimum k ’values for maximum resolution are between roughly 1 and 4 for toft, values of 0.2 or greater. The relationship of eq 7-9 to the separation of oligonucleotides with micelles and added metal ions will be discussed in a later section of the paper after the mechanism in this case has been described. Bases and Nucleosides. In order to separate the uncharged bases and nucleosides a t neutral pH, advantage has been taken of solute partition into the interior of a micelle. An anionic surfactant, sodium dodecyl sulfate (SDS), was selected to form the micelles, since the charge on the fused silica capillary walls is also negative and electrostatic adsorption of the surfactant to the walls will then not occur. Figure 1A shows the capillary electrophoretic separation of four bases in 0.1 M SDS, pH 7.0, under a potential of 14 kV and a current of 50 FA. The peak sharpness and rapid separation are especially to be noted. The peak a t 5.2 min is equal to toand arises from the injected solvent peak. The elution order is based on increasing hydrophobicity of the solutes. Ura (see end of paper for abbreviations), the most hydrophilic species, has the shortest migration time, while Ade, the most hydrophobic species, has the longest. Since the interior of the SDS micelle is hydrophobic, the more hydrophobic the species the greater will be its partition into the micelle (i.e,, k’will be higher the more hydrophobic the solute). Since the negatively charged micelle moves through the capillary more slowly than the bulk osmotic flow, the slower migration of the more hydrophobic species is understood. Figure 1B shows the separation of nucleosides, and again elution is in the order of increasing solute hydrophobicity; however, the ribose ring makes all these species more hydrophilic than their corresponding bases, and the elution conditions are thus slightly different relative to Figure 1A. The SDS concentration is doubled to 0.2 M in orcler to produce a higher number of micelles (thus increasing k’through an increase in V,, eq 8 and the applied potential is decreased to 9.5 kV (40 FA) in order to prevent overheating of the capillary. Figure 1C shows an example of the mixture of nine bases and nucleosides. As noted above, under common conditions the nucleosides elute earlier than their corresponding bases. In this case, the applied voltage is 5.9 kV which yields a current of 30 FA. Note the increased broadness of the peaks in Figure lC, relative to Figure 1A,B. An increase in voltage decreased
ANALYTICAL CHEMISTRY, VOL. 59, NO. 7, APRIL 1, 1987
1023
A C
51
.__a_ I
1
2
4
I
6
I
I
a
5
T i me / m i n
I
10 15 Timermi n
I
16
20
20
24
28
32
36
Timelmin
Flgure 1. (A) Separation of bases: (1) Ura, (2) Cyt, (3) Thy, (4) Ade (see end of paper for abbreviations); buffer, 0.025 M sodium tetraborate, 0.05 M sodium dihydrogen phosphate, 0.1 M SDS, pH 7; capillary, 650 mm X 0.5 mm i.d., effective length 500 mm; applied voltage, 14 kV, 50 PA; detection wavelength, 210 nm. The system was air controlled at 35 OC. (B) Separation of nucleosides: (5)Urd, (6) Cyd, (7) dThd, (8) Guo, (9)Ado (see end of paper for abbreviations). Condttbns are as given In part A, except 0.2 M SDS and 9.5 kV applied voltage, 40 pA. (C) Separation of bases and nucleosides. Conditions are as given in part A, except 0.3 M SDS and 5.9 kV applied voltage, 30 pA.
Table I. Enthalpy, Entropy, and Distribution Coefficient in Micellar Solubilization of Bases and Nucleosides with SDS Solution"
0
20
40
60
I /PA
80
Flgure 2. Dependence of the solute capacity factor, k', on the current. Buffer and capillary conditions are given in Flgure 18. The numbers
of the plot correspond to the solutes in Figure 1. separation time and sharpened peaks; however, full separation was not achieved. The increased current accompanying the increase in voltage could have caused a temperature rise due to increased Joule heating (the capillary walls were only air-cooled in this case), and this could have affected the partition process (25). In order to examin? whether this explanation is valid, we decided to explore k'as a function of current generated in the capillary. Figure 2 shows the effect of current on the &'of the bases and nucleosides using 0.2 M SDS. The capacity factor ,&'was determined from eq 10 which is simply a rearrangement of eq 7
It is seen in Figure 2 that &'decreases linearly with current, but the slopes differ and changes in selectivity are thus observed. These changes are a consequence of the Joule heating effects on the solute distribution coefficient K.
Wo/kJ mol-'
AS"/J K-'
solute
mol-'
K(40 "C)
Ura Urd Cyd CYt dThd Guo Thy Ado Ade
-15.5 -12.5 -16.4 -21.5 -15.8 -21.6 -16.4 -23.9 -23.9
-34 -25 -35 -50 -31 -48 -3 1 -54 -53
6.4 6.4 7.; 9.6 11.2 12.6 12.6 15.3 16.3
"0.2 M, pH 7. Calculated from distribution coefficients measured at 41,48, 57, and 61 "C.
The enthalpies and entropies of micelle partition for the nine solutes were measured in a manner previously described (15),and the results are shown in Table I for an estimated temperature of 41 "C (-30 FA under the buffer conditions of Figure 3). It can seen that Ura and Urd as well as Guo and Thy have similar K values at this current; however, the AH" values are sufficiently different that a t other currents (temperatures) separation can be achieved. Thus, temperature can be used as a means of selectivity control in the capillary electrophoretic separation of neutral species with micelles. The results of Table I further emphasize that reproducibility in separation and migration times requires good temperature control. We have recently been using a liquid thermostated system for precise temperature control (see later). In Figure lC, the current of 30 FA represents less Joule heating than in Figure 2, since the solution resistance created by the 0.3 M SDS solution is less than that of 0.2 M SDS. Evidently, the temperature is sufficiently low to achieve separation (i.e., corresponding to a temperature of -20 pA in Figure 2). As we have already noted, increasing voltage with the conditions of Figure 1C caused overlap of peaks in spite of the increased band sharpness. This result can be understood with reference to Figure 2. An increase in voltage any thus current will increase temperature and likely alter the k '
1024
ANALYTICAL CHEMISTRY, VOL. 59, NO. 7, APRIL 1, 1987
,T 6
,/'
A
B
I
5
7 -3------Q
0
6l OT
i
9
9 2
I1
Flgure 3. Dependence of retention time, t,, on pH. Buffer used is the same as that given in Figure 1B except pH adjusted. The numbers on the plot correspond to the solutes in Figure 1. The species are identified under the abbreviation list at the end of the paper.
values into a region of overlap (e.g., equivalent to -30 pA in Figure 2). Figure 3 demonstrates the dependence of migration time of the bases and nucleosides on the p H of the buffer in the range of pH 7-9. The considerable increase in migration time for Urd (no. 5 ) , Guo (no. 8), and Ura (no. 1)with increasing pH is due to the partial ionization of the solutes to negatively charged species. The pK, values for deprotonated forms of these compounds are 9.2, 9.2, and 9.5 for Urd, Guo, and Ura, respectively. These results suggest that the electrophoretic effect on partially ionized solutes considerably reduces the migration velocities of these solutes. Cyd (no. 6) and Ado (no. 9) also increase migration times with increasing pH; however, charge variation cannot account for the changes since the pKA for formation of a negative charged species (ribose ionization) is 12.5. The reason for the behavior of Cyd and Ado in Figure 3 is not clear at present. All the other solutes are not affected in the pH range examined, and their migration time remains constant. Figure 3 demonstrates that pH can be used to affect selectivity in the separation of these species. Oligonucleotides. Oligonucleotides are negatively charged a t neutral pH, and they can thus be studied directly by capillary electrophoresis. Moreover, they will be electrostatically repelled from the negatively charged fused silica surface so that adsorption which could cause band broadening is not anticipated. Figure 4A shows the separation of a polythymidine mixture a t pH 7 and 7 M urea (to prevent aggregation of complementary oligonucleotides (13))with an applied potential of 20 kV (10 PA). In this example, as well as all others with oligonucleotides, the capillary was thermostated at 25 "C with carbon tetrachloride. Separation is observed up to 10 bases; however, overlap occurs for the mixture of 12-18 bases. The reason for this overlap is due to the well-known small differences in electrophoretic mobility for oligonucleotides as the base number increases (13). As a second step, 0.05 M SDS was added to the buffer to create conditions similar to those employed to separate the bases and nucleosides. Figure 4B shows the separation of the same polythymidine mixture with SDS micelles present. Some resolution is still observed; however, it is poorer than that in Figure 4A. The oligonucleotides also elute earlier with the SDS present, and this appears to be a consequence of a slightly higher electroosmotic velocity (ueo)in Figure 4B, relative to Figure 4A (tois less with SDS present in the buffer). In Figure 4, parts A and B, the spacing between bands is minimal. While some improvement in separation can be
'c, Lu 0
Flgwe 4. (A) Separation of polythymidines [ 1, solvent; 2, d(pT)*; 3, d(PT),; 49 d(PT)i 5, d(PT),; 6, d(PT),o; 7, d(pT),,-d(pT)1*; pT = polythymidine]: buffer, 7 M urea, 5 mM Tris, 5 mM Na2HP0,, pH 7; capillary, 650 mm X 0.05 mm i.d., effective length 450 mm, applied voltage 20 kV (10 PA), detection wavelength 260 nm, thermostated at 25 'C. (E) Ail conditions are the same as those given in part A except addition of 50 mM SDS to the buffer.
expected by manipulation of the physical characteristics of the system (e.g., voltage, capillary diameter), the resolving power would probably remain inadequate for complex mixtures of oligonucleotides up to -10-12 bases. Accordingly, we searched for chemical means to improve selectivity and band spacing. Metals, e.g., Cu(II), Zn(II), and Mg(II), are known to complex with oligonucleotides, with either the phosphate groups and/or the bases (18,19). We therefore explored the possibility of employing selective metal-oligonucleotide complexation to manipulate separation of these species. We first examined the addition of 0.3 mM Cu(I1) and 5 mM Mg(I1) to the trisphosphate buffer a t pH 7 (SDS was not present). For the Cu(I1) buffer the electroosmotic velocity decreased by a factor of 2 and that of Mg(I1) by a factor of 1.5, relative to when no metal was present. These metals would be expected to be adsorbed to the walls of the fused silica at neutral pH (20),thus lowering the zeta potential of the capillary walls. Equation 1 shows that a lower zeta potential leads to a slower electroosmotic velocity. In addition, separation of oligonucleotides with added metal led to significantly broadened peaks, undoubtedly due to the adsorption of the solutes onto the metal sites at the surface of the fused silica. An alternative approach was thus necessary to employ metal ions in the separation of oligonucleotides. We next added 50 mM SDS to the metal-buffer solution and found a significant increase in the u,, values. Moreover, the polythymidines are well resolved from one another with relatively sharp elution peaks, as seen in Figure 5. A dramatic improvement is observed, relative to Figure 4. A comparison of Figure 5A and Figure 5B also clearly shows better band spacing in the case of Cu(I1) (resolution up to 18 bases of polythymidine), although the peaks appear relatively sharper in the case of Mg(I1). The wider time window in the case of Cu(I1) is related to stronger oligonucleotide-metal binding for this metal relative to Mg(I1) (19). The smaller u, (or larger to)value for Cu(I1) suggests a greater amount of metal ad-
ANALYTICAL CHEMISTRY, VOL. 59, NO. 7, APRIL 1, 1987
1025 I
B
7
5
/I
L
ri
LLL
0
10
5
mc
to
A 4
I
4
I
0.1
1
I
I
I
I
0.2
03
0.4
0.5
0.6
3 I
Conc. o f Cu++ (mM) 0
10
20
30
40
50
Figure 6, Plot of k’,,,c of the SDS micelle and t o vs. Cu(I1) concentration added to the buffer. All other conditions are given in Figure 48.
Figure 5. (A) Separation of polythymidlne mixture. 1-6, as in Figure 4; 7, d(pT),,; 8, d(pT),,-d(pT),,. All conditions are the same as those given in Figure 48 except addition of 5 mM Mg(I1) to the buffer. (8) Separation of polythymidine mixture: 1-7, as in Figure 5A; 8, d(pT),,; 9, d(pT),,; 10, qpT),,; 11, ~(PT),,; 12, (YpT),,; 13, d(pV18.. All conditions are the same as those given in Figure 46 except addition of 0.3 mM Cu(I1) to the buffer.
sorption on the fused silica walls for this metal relative to Mg(I1). Cu(I1) is known to bind more strongly to silica than does Mg(I1) at neutral pH (21). The broadening in Figure 5B may be due to slight adsorption of the oligonucleotide to the Cu(I1)-fused silica surface. Nevertheless, the use of metals and SDS micelles is clearly beneficial in separation of oligonucleotides. Consider next the mechanism of migration of the solutes in Figure 5A,B. Metal ions are known to adsorb electrostatically to the surface of anionic micelles (16). (50 mM SDS is a concentration well above the critical micelle concentration (cmc), even with metal present in solution.) Undoubtedly, the micelles provide a competitive surface to the negatively charged fused silica capillary walls for the metal ions. As a consequence, less metal is adsorbed to the walls, thus minimizing solute adsorption and band broadening. The increased zeta potential, as reflected in the lower tovalues upon the addition of SDS to the metal-buffer solution, is consistent with less metal adsorbed on the surface of the walls. As further evidence for the adsorption of metal ion to the surface of the micelle, we measured toand kLC(capacity factor ) a 50 mM SDS solution as a of the micelle = (tmc- t o ) / t oof function of copper concentration, and the results are shown in Figure 6. As expected, toincreases with added Cu(II), since even with SDS present, a small amount of metal will adsorb to the walls and decrease the zeta potential of that surface. Concurrently, k kcdecreases with Cu(I1) added to the buffer system. As the metal is adsorbed to the surface of the micelle, the zeta potential of the micelle will be reduced due to charge neutralization, and the micelle will then migrate faster in the capillary. Figure 7 presents a schematic representation of the likely
Flgure 7. Schematic illustration of retention mechanism with SDS micelles and metal ions.
mechanism of separation. Each oligonucleotide will possess a given electrophoretic velocity, uep, dependent on the electrophoretic mobility of the species, pep,and the electrophoretic conditions, eq 4. This mobility could be altered by equilibrium of the solute with free metal present in the solution; however, since the amount of free metal is expected to be low in the presence of SDS micelles, we shall to a first approximation ignore this effect. In addition, retardation may occur by oligonucleotide adsorption on metal sites at the fused silica walls. For purposes of this discussion, we shall also ignore this effect. A major component of the migration mechanism will be oligonucleotide complexation with metal ions attached to the surface of the micelles. As in the case of neutral species partitioning within the interior of the micelle, the attachment of the nucleotide to the micelle will result in a reduction in the migration velocity of the solute. The time window over which separation is possible will thus be significantly enhanced, leading to markedly improved resolution, as observed in Figure 5. Kinetically, this association-dissociation would be expected to be quite rapid (22),and the sharp peaks of Figure 5 confirm this expectation. The principles developed for neutral species with micellar capillary electrophoresis ought to apply for the mechanism
1026
ANALYTICAL CHEMISTRY, VOL. 59, NO. 7, APRIL 1, 1987
-
.
j
l
l
I
l
l
I
I
10 I
l
I I 1 i I
20
1 1 I I 1 1 I I
30
40
I 1 1 l l I I I l I 1 1 1
Figure 9, Separation of a mixture of 18 oligonucleotides, each with 8 bases. Separation conditions are the same as those given in Figure 8C except that the length of the tube was 850 mm and 22 kV (10bA) was employed.
1
A
< Cu(I1). The zeta potential is inversely proportional to to, eq 1, and the above order is easily understood in terms of the extent of adsorption of the metals on the fused silica surface (21). 1 1 1 1 I I I I 1 1 I I I I l 1 1 l l l l , , , I 1 / I I I I I t I / / I I I / I I ,1 1 l l l I I 1 1 1 1 ~
0
20
Figure 8. Effect of metal ions on the separation of oligonucleotides, each with eight bases: (1) water; (2) CATCGATG; (3) AACGCGTT; (4) GGGATCCC; (5) AAAGCTTT; (6) CGGGCCCG (7) CGCCGGCG. (A) Buffer: 7 M urea, 20 mM Tris, 5 mM Na2HP0,, 50 mM SDS. All other conditions are given in Figure 4A. (B) Addition of 3 mM Mg(II) to the buffer. (C) Addition of 3 mM Zn(I1) to the buffer. (D) Addition of 3 mM Cu(I1) to the buffer.
of Figure 7. The capacity factor k ’ (eq 10) will now represent the binding of oligonucleotide to the micelle. The value of i fcan be expressed as
where t’* is the migration time of the solute in the presence of the metal-micelle system, and K, is the complexation constant of the oligonucleotide with the metal-micelle surface. In this equation, t,, the migration time without SDS or metal present, is used instead of t o (cf. eq lo), since the oligonucleotide will migrate at a different velocity than ueo; see eq 6. Separation will be related in large measure to differences in K, values for the various oligonucleotides. Manipulation of K , via changes in metal and surfactant as well as the buffer conditions, ought to provide separation differences. It is known that Mg(I1) coordinates with the phosphate groups, while Cu(I1) interacts strongly with the bases and only weakly with the phosphate groups (18, 19). Based on the results of Figure 5B, Cu(I1) would appear a good choice of metal, given its strong binding to oligonucleotides. However, Cu(I1) is UV active and a possible alternative metal could be Zn(I1) which is UV transparent in the range of oligonucleotide detection. Zn(I1) is intermediate in oligonucleotide binding strength between Mg(I1) and Cu(I1). Figure 8 shows the separation of a mixture of six oligonucleotides of eight bases, each with a different sequence, as a function of metal ion (3 mM) added to the SDS micellebuffer solution. We first note that for the same concentration of metal, the tovalue is in the order no metal < Mg(I1) < Zn(I1)
Separation appears best in the case of Zn(II), Figure 8C. More overlap of peaks can be seen for Mg(1I) for which weak K , values exist. In the case of Cu(II), good band spacing is observed; however, the peaks are significantly broader. This result is probably due to increased oligonucleotide adsorption on the metal-silica wall surface. Note that 3 mM Cu(I1) is used in this example, a 10-fold increase in concentration to that employed in Figure 5. Lower concentrations of Cu(I1) may yield an appropriate compromise of band separation and width; however, this was not investigated. In the case of Zn(II), sharp bands and full separation are observed. The power of this approach can be seen in Figure 9 which illustrates the resolution in less than 30 min of 14 out of 18 oligonucleotides of 8 bases, each with a different sequence. CONCLUSIONS This paper has shown the further value of hgh-performance micelle-capillary electrophoresis. As in a previous paper (15), this work illustrates the approach for separation of neutral species via partition within the micelle. In addition, the results presented here demonstrate that the surface of the micelle can also be used to enhance separation of charged species. Metal ions are electrostatically attracted to the surface of the micelle where they can be selectively complexed with appropriate solutes. This approach has three advantages. First, it has the potential to broaden the time window over which separation can occur (thus increasing peak capacity) (15). Second, complexation differences in molecules can be employed to manipulate selectivity. Third, the kinetics of the association-dissociation processes appear sufficiently rapid to minimize band broadening, even for the demanding procedure of capillary electrophoresis. ACKNOWLEDGMENT J. M. Smith thanks M. Stempowski and L. Kizilay for technical assistance. S. Terabe thanks H. Utsumi for technical assistance. Bases 1. Ura, uracil
NOMENCLATURE
Anal. Chem. 1987, 59, 1027-1031
2. Cyt, cytosine 3. Thy, thymine 4. Ade, adenine
Nucleosides 5. Urd, uridine 6. Cyd, cytidine 7. dThd, deoxythymidine 8. Guo guanosine 9. Ado, adenosine 10. dpT, deoxypolythymidine LITERATURE CITED Jorgenson, J. W.; Lukas, K. D. Science 1983, 222, 266-272. Hjerten, S.;Zhu, D. M. J . Chromatogr. 1985, 346, 265-270. Tsuda, T.; Nomua. K.; Nakagawa, G. J . Chromatogr. 1982, 248, 241-247. Terabe, S.;Otsuka, K.; Ichikawa, K.; Tsuchiya, A.; Ando, T. Anal. Chem. 1984, 56, 111-113. Lauer, H. H.; McManigill, D. Anal. Chem. 1988, 58, 166-170. Gassmann, E.: Kno, J. E.;Zare. R. N. Science 1985, 230, 813-815. Hjerten, S. J . Chromatogr. 1985, 347, 191-198. Aveyard, R.; Haydon, D. A. An Introduction to the Principles of Surface Chemistry; Cambridge University Press: Cambridge, 1973; Chapter 2. Everaerts, F. M.; Mikkers, F. E. p.; Verheggen, Th. p. E. M.; Vacik, J. In Chromatography,Part A ; Heftmann, E., Ed.; Eisevier: Amsterdam, 1983; Chapter 9.
1027
Giddings, J. C. S e p . Sci. 1969, 4 , 181-189. Dizaroglu, M.; Hermes, W. J . Chromatogr. 1979, 171, 321-330. Brown, P. R.; Grushka, E. Anal. Chem. 1980, 52, 1210-1215. Gel fiectrophoresis of Nucleic Acids: A Practical Approach; Rickwood, D., Hames, B. D., Eds.; IRL Press: Washington, DC, 1983. Tsuda, T.; Hakagawa, G.; Tato, M.; Yagi, K. J . Appi. Biochem. 1983, 5 , 330-336. Terabe, S.; Otsuka, K.; Ando, T. Anal. Chem. 1965, 5 7 , 834-841. Oko, M. U.; Venabie, R. L. J . Coiioid Interface Sci. 1971, 35, 53-59. Hunter, R. J. Zeta Potential in Colioid Science; Academic: London, 1981; Chapter 3. Scheller, K. H.; Hofstetter, F.; Mlchell, P. R.; Prijs, B.; Sigel, H. J . Am. Chem. SOC. 1981, 103, 247-260. Metal Ions in Bioiogicai Systems; Sigei, H., Ed.; Marcel Dekker: New York, 1979; Voi. 9. (20) Iler, R. K. The Chemistry of Silica ; Wiley: New York, 1978. (21) Wakatsuki, T.; Furukawa, H.; Kawaguchi, K. Soil Sci. Piant Nutr. (Tokyo) 1974, 2 0 , 353. (22) Burgess, J. Metal Ions in Solutions; Halsted Press: New York, 1978.
RECEIVED for review September 23,1986. Accepted December 18, 1986. B. L. Karger gratefully acknowledges support by the James L. Waters Chair in Analytical Chemistry, the National Science Foundation, and Dow Chemical Company. J. A- Smith gratefully acknowledges support by I'bXhst Aktiengesellschaft (FRG). This is Contribution No. 299 from ~and Materials ~ l the ^ . Barnett Institute of Chemical ~ Science.
Determination of Organic Compounds Leached from Municipal Incinerator Fly Ash by Water at Different pH Levels F. W. Karasek,* G . M. Charbonneau, G . J. Reuel, and H. Y. Tong Department of Chemistry, University of Waterloo, Waterloo, Ontario, Canada N2L 3G1
This study concerns the possiblilty of organlc compounds entering the environment through the leaching of municipal Incinerator fly ash wtth water. A Soxhlet extractlon of fly ash with water, followed by a benrene/water solvent extractlon was used to Isolate organlc compounds. The pH of the extracting llquld was varled (pH 4, 7, 10) and both the type and amount of compounds extracted dlffered. Many organlc compounds Including polychlorinated dibenzo-p -dloxlns, polychlorinated dlbenzofurans, polycyclic aromatic compounds, phenols, and hydrocarbons were found in the water extracts. Gas chromatographylfiame ionization detectton, gas chromatography/electron capture detectlon, and gas chromatography/mass spectrometry uslng electron Impact, posltlve and negative Ion chemical lonlzatlon techniques were used for compound identlficatlon and quantltatlon.
Fly ash, a fine particulate effluent from municipal incinerators, is the major byproduct produced from the burning of municipal waste. Approximately 35 000 tons of fly ash are produced for each million tons of waste incinerated (1). Considering the amount of municipal garbage incinerated in cities worldwide, the quantity of fly ash is significant. More than 600 organic compounds are known to be adsorbed on these particulates, 200 of which have been identified (2). Although electrostatic precipitators are used to remove the fly ash from the flue gas, approximately 1-2% of the produced
fly ash escapes to the atmosphere (I). Most studies have been carried out to determine the effect of the escaping fly ash (34). Little if any investigation has been done regarding the potential harmful effects of the disposal of the majority of the fly ash in landfill sites. Much of the fly ash buried is constantly exposed to rainwater, groundwater, and possibly wastewater as well. This exposure raises an important question as to whether organic pollutants in the fly ash could be removed by water and eventually contaminate the environment. Little if any investigation has been done regarding this possibility. Soxhlet extraction of the fly ash was used to resemble leaching conditions in which fly ash comes into contact with water and to provide a concentrated extract from which organic compounds could easily be isolated. Fresh solvent is passed through the fly ash during each cycle, and an exhaustive extraction would show the extent of leaching in the most serious case, where fly ash is constantly exposed to an aqueous environment. Since the pH of water greatly affects the type of compounds extracted in a typical solvent extraction, the pH of the extracting water was also altered. By the addition of formic acid and 2-methylethylamine, extractions were carried out at a pH of 4 and 10 in addition to pH 7. Organic compounds were then transferred to an organic solvent. A gas chromatographic analytical method consisting of five detection techniques was used for the analysis of organic compounds in the water extracts; flame ionization detection (GC/FID); electron capture detection (GC/ECD); mass
0003-2700/87/0359-1027$01.50/0 0 1987 American Chemical Society
~
