INSTRUMENTATION
Liquid Chromatography with Pulsed Electrochemical Detection Dennis C. Johnson and William R. LaCourse Department of Chemistry Iowa State University Ames, IA 50011
The development of detection methods for aliphatic compounds in LC has rep resented a challenge of major analyti cal proportion. Significant advances have occurred in the use of preinjection and postcolumn chemical derivatizations to produce photometrically and electrochemically active adducts (1, 2). Historically, however, researchers have agreed that aliphatic compounds are not amenable to amperometric detec tion (3). We believe that the simplicity of sensitive direct detection in LC will always be preferred, whenever avail able. In this article, we will describe the theory and use of multistep potential waveforms (E-t) for direct, sensitive, and reproducible detection of alcohols, glycols, carbohydrates, alkanolamines, amino acids, and sulfur compounds at Au and Pt electrodes. Numerous aromatic compounds are detected easily by anodic reactions at a constant (dc) applied potential at solid electrodes—for example, Au, Pt, and various forms of carbon (3, 4). Signifi cant examples include phenol, halogenated phenols, aminophenols, cate cholamines, and other metabolic amines. In contrast, however, the ma jority of aliphatic alcohols and amines is not consistently observed to be electroactive under conditions of ampero metric detection at constant (dc) ap plied potentials. We have concluded that this differ ence in reactivity results from the lack of conjugated bonding in the anodic re 0003-2700/90/0362-589A/$02.50/0 © 1990 American Chemical Society
action mechanisms for aliphatic com pounds. Free-radical products from ox idations of aromatic molecules can be stabilized by ττ-resonance; hence, the activation barrier for the reaction is de creased. A similar mechanism for stabi lization of aliphatic free radicals is ab sent and, consequently, oxidation rates are normally very low, even though an odic reactivity is predicted based on thermodynamic data. The absence of 7r-bonding in aliphatic compounds also results in the absence of sensitivity for photometric (UV-vis) detection of these compounds. The activation barrier for oxidation of aliphatic compounds can be de creased at noble-metal electrodes (e.g., Au and Pt) with partially unsaturated surface d-orbitals that can adsorb and thereby stabilize free-radical interme diate oxidation products. However, a serious consequence of strong adsorp tion can be the fouling of these elec trodes by accumulated detection prod ucts (5-8). Hence, the historical con sensus of nonreactivity for aliphatic compounds at Au and Pt electrodes is the result, in many instances, of a high but relatively short-lived catalytic ac tivity for these electrodes in t h e "clean" state. Regeneration of electrode activity Adsorbed hydrocarbons can be oxidatively desorbed quite efficiently from these electrodes by application of a large positive potential excursion, which causes the formation of surface oxide (5-9). The intermediate products in the oxide formation (AuOH and PtOH) are reactive in the mechanism of oxygen transfer from H2O to the oxi dation products. However, the final stable oxides (AuO and PtO) are quite
at Gold and Platinum Electrodes inert and must be cathodically dis solved by a negative potential excur sion to restore the native reactivity of the clean metal surfaces. Some organic compounds are not electroactive at the oxide-free noble-metal surfaces but are adsorbed at these surfaces. These com pounds also can be oxidatively desorbed simultaneously with the oxide formation process to give useful anodic signals. The application of large potential ex cursions at noble-metal electrodes has long been known to result in the prepa ration of reproducibly clean and reac tive electrode surfaces. Virtually every publication on voltammetric data for noble-metal electrodes briefly de scribes a protocol for electrode pretreatment that was applied to maintain electrode activity and give reproduc ible data. Most commonly, pretreatment includes the application of re peated cyclic potential scans or alter nated positive and negative potential pulses. Such electrochemical treat ment for maintaining the highest activ ity of P t electrodes was reported by
ANALYTICAL CHEMISTRY, VOL. 62, NO. 10, MAY 15, 1990 · 589 A
INSTRUMENTATION Hammett in 1924 (10) for the study of H 2 oxidation and by Armstrong et al. in 1934 for studies of O2 reduction (11). More recently, Clark et al. (12) re ported greater reproducibility for ano dic oxidation of ethylene at P t when cleaning pulses were applied, and MacDonald and Duke (13) offered a similar report relating to the detection of p-aminophenol. Comparable benefits of pulsed potential cleaning have been obtained also for detection of inorganic species at noble metals, as reported by Stulik and Hora (14) for the detection of Fe(III) and Cu(II) at Pt. The in creased sensitivity and reproducibility from the application of potential pulses at carbon electrodes also have been claimed by several workers, including Fleet and Little (15), van Rooijen and Poppe (16), Ewing et al. (17), Berger (18), and Tenygl (19). Concepts of pulsed electrocatalytic detection Faradaic processes that benefit from interaction of the electrode surface within the reaction mechanism are de scribed as "electrocatalytic." Based on this discussion, two modes of anodic electrocatalytic detection are obvious at noble-metal electrodes in conjunc tion with the application of potential pulses to achieve anodic cleaning and cathodic reactivation of electrodes. Mode I: Direct detection at oxidefree surfaces. There is very little or no concurrent formation of oxide in this detection mode. The baseline signal originates primarily from double-layer charging and decays quickly to a virtu al zero value. Simple alcohols, glycols, so-called sugar alcohols, monosaccha rides, and oligosaccharides are detect ed by Mode I at Au electrodes in alka line solutions and at Pt electrodes in alkaline and acidic solutions (20-30). Mode II: Direct oxide-catalyzed detection. Oxidation of adsorbed analyte is the primary contributor to the analytical signal in this detection mode; however, simultaneous catalytic oxidation of analyte in the diffusion layer is not excluded. The background signal is large, originating from anodic formation of surface oxide. Aliphatic amines and amino acids (31-33) as well as numerous sulfur compounds (34-37) are detected by Mode II at Au and P t electrodes in alkaline solutions. A consequence of the electrocatalytic basis of detection is that the amperometric response of various members within a class of compounds is con trolled primarily by the dependence of the catalytic surface state on the elec trode potential rather than by the re dox potentials (E°) of the reactants. Hence, there is little hope for voltam
metric resolution of complex mixtures, and electrocatalytic detection will be most useful when coupled to LC. Au electrodes have become the most significant of the noble metals for pulsed electrochemical detection. Fig ure 1 illustrates various potential re gions of response by Modes I and II for Au in 0.1 M NaOH superimposed on the residual i-E curve obtained during a cyclic potential scan with respect to a standard calomel electrode (SCE) ref erence. Surface oxide is formed during the positive scan for Ε > +0.2 V (Wave A), and solvent breakdown with O2 evolution occurs for Ε > ~+0.6 V (Wave B). During the subsequent neg ative scan, O2 evolution and oxide for mation cease for Ε < +0.6 V and ca thodic dissolution of the surface oxide produces the cathodic peak in the re gion +0.2 V to - 0 . 1 V (Wave C). If present, dissolved O2 (indicated by a dotted line) is cathodically detected at Ε < — 0 . 1 V (Wave D). All aldehydes, including the so-
called reducing sugars, are anodically detected during a positive potential change at the oxide-free surface in the region — 0 . 6 V to +0.2 V (Mode I). Large anodic signals are obtained for alcohols, polyalcohols, and nonreducing sugars in the region ~—0.3 V to +0.2 V (Mode I) with much less intense signals for some compounds from +0.2 V to +0.6 V (Mode II). Amines and sulfur-containing compounds, for which a nonbonded electron pair re sides on the N- and S-atoms, are ad sorbed at oxide-free Au surfaces for Ε < — 0 . 1 V and can be anodically detected by oxide-catalyzed reactions during the positive scan for Ε > ~ + 0 . 1 V (Mode II). Detections at Ε > ~+0.6 V are not recommended because of the evolution of O2. Detection strategies Pulsed electrochemical detection. Amperometric detection based on the electrocatalytic mechanisms under dis cussion can be accomplished automati-
Figure 1. Residual voltammetric response (i-E) for an Au rotated disk electrode in 0.1 M NaOH. Conditions: 200 rev m i n - 1 rotation, 100 mV s~1 potential scan. Waves (/—£): A, oxide formation (positive scan); B, 0 2 evolution; C, oxide reduction (negative scan); D, 0 2 reduction. Anodic response (positive scan): I, aldehydes and reducing sugars; II, alcohols and nonreducing sugars; III, amines and amino acids; IV, sulfur compounds.
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Figure 2. Potential-time (E-t) waveforms. Processes: £,, anodic detection; £2, oxidative cleaning; £3, cathodic reactivation. Waveforms: (a) pulsed amperometric detection (PAD) with a short current sampling period (e.g., 16.7 ms), (b) PAD with a long current integration period (e.g., 200 ms), and (c) integrated PAD with a long integration period (e.g., 200 ms).
cally by the multistep potential wave form illustrated in Figure 2a (20-30). The detection potential (Ει) is chosen to be appropriate for the desired detec tion mechanism (see Figure 1), and the faradaic signal can be sampled during a short time (e.g., 16.7 ms) after a delay of id near the end of the detection peri od (ti). Typical values of ii are in the 100-400-ms range. Following the de tection process, the electrode surface is oxidatively cleaned by a positive step to £2 (h = 50-200 ms) and then cathodically reactivated by the large negative step to E3 (i 3 = 100-400 ms) prior to the next detection cycle. Typically, the waveforms are executed at a frequency of ~ l - 2 Hz, which generally is appro priate for detection in LC. Amperome tric detection under the control of a multistep waveform commonly has been called pulsed amperometric de tection (PAD). The origin of detection peaks in LC using PAD based on Mode I is illustrat ed in Figure 3a by generic chronoamperometric (i-t) response curves following the step from Es to E\ at an oxide-free electrode (30). The residual response from double-layer charging (curve A) decays very quickly, and the baseline signal in LC-PAD is very small for id > ~50 ms. Curve Β in Figure 3a represents the i-t response for the presence of analyte, and the arrow rep resents the corresponding signal ex pected for LC-PAD with the indicated value of ti in the waveform. The origins of detection peaks in LCPAD based on Mode II are illustrated
in Figure 3b (30). Here, the residual i-t response (curve A) corresponds to the formation of surface oxide, which de cays much more slowly than the cur rent from double-layer charging. Base line signals for Mode II typically have a nonzero value, as illustrated in Figure 3b for two values of id. The i-t response (curve Β in Figure 3b) corresponds to the presence of adsorbed analyte, and arrows are shown for the two values of
id to represent the corresponding LCPAD response. For small id, "negative" peaks can be obtained because of initial inhibition of the oxide formation pro cess by the adsorbed analyte. For larger values of id, "positive" peaks are ob tained when sufficient oxide has been produced to catalyze the anodic reac tion of the adsorbate. If an intermedi ate value of id is chosen, a detection peak might not be obtained. Choosing id > ~150 ms usually is sufficient to assure "positive" LC-PAD peaks based on Mode II detection. Sampling of electrode response in PAD. The signal-to-noise ratio (S/N) for measurements of transient ampero metric signals is influenced by the in strumental strategy used for sampling the electrode current. A major noise component of the chronoamperometric signal is sinusoidal and correlated with the 60-Hz line frequency. Hence a com mon strategy for current sampling in PAD involves some form of signal aver aging over the period of one 60-Hz os cillation (i.e., 16.7 ms). Accordingly, there is no contribution to signal strength from the 60-Hz noise. Actual ly, the time integral of a 60-Hz sinusoi dal noise signal is zero for integration over any integral number (m) of 16.7ms periods (38). Yet the analytical sig nal strength can be increased signifi cantly for m » 1. As an example, if the analytical signal is a constant value throughout the period m · 16.7 ms, the S/N will be increased by the factor m. Typically, m = 12 and the integration period (i;) is 200 ms. This longer sam-
Figure 3. Chronoamperometric response {i-t) following a potential step from E3 to E\ in the PAD waveform to illustrate the origins of chromatographic baseline and peak signals in LC-PAD. Detection: (a) Mode I and (b) Mode II. Curves: (A) Background response in the absence of analyte and (B) response in the presence of analyte. Note that delay after step to Ei is indicated by tj, LC baselines are indicated by the labeled dashed lines, and LC-PAD peak signals are indicated by arrows.
ANALYTICAL CHEMISTRY, VOL. 62, NO. 10, MAY 15, 1990 · 591 A
INSTRUMENTATION pling period for PAD is illustrated by the waveform in Figure 2b. Because the signal output for an integrated amperometric response has units of cou lombs, the corresponding technique was originally called pulsed coulometric detection (38). Integrated PAD with a potential sweep. As discussed for Figure 3b, a large baseline signal is encountered in LC-PAD for the oxide-catalyzed detec tions of amino acids and sulfur com pounds (Mode II). Furthermore, the large baseline current is frequently ob served to drift to more anodic values, especially for new or freshly polished electrodes. This drift occurs because of slow growth in the true electrode sur face area that is attributable to surface reconstruction caused by the oxide onoff cycles in the applied multistep waveforms. Baseline offset and drift can be sig nificantly diminished by use of the waveform in Figure 2c (39). Here, the electrode current is integrated throughout a rapid cyclic scan of the detection potential CEi) within a pulsed waveform. The potential scan proceeds into (positive scan) and back out of (negative scan) the region of the oxide-catalyzed reaction for detection by Mode II. The anodic charge for ox ide formed on the positive sweep dur ing the detection period tends to be compensated by the corresponding cathodic charge (opposite polarity) for dissolution of the oxide on the negative sweep. Hence the "background" signal on the electronic integrator at the end of the detection period can be virtually zero and is relatively unaffected by the gradual change of electrode area. The detection procedure based on the waveform in Figure 2c was originally called potential scan pulsed coulometric detection (39); however, we now prefer the name integrated pulsed amperometric detection (IPAD).
taining dissolved 0 2 . The purging of solvents with He and use of 0 2 -impermeabie tubing can result in virtually Cvfree conditions that greatly relax the constraints of these two criteria. The advantage of IPAD compared with PAD relates to minimization of baseline drift for oxide-catalyzed de tections (Mode II). Comparisons of IPAD with PAD for carbohydrates at the oxide-free surfaces (Mode I) have indicated virtually no significant dif ference in detectability for the two techniques, and we recommend contin ued use of PAD (i; = 200 ms) for carbo hydrate and alcohol detection. An additional consideration in IPAD relates to the electrochemical revers ibility of the detection reaction. Clear ly, the anodic signal is expected to be at a maximum when there is no cathodic contribution to the net current integral from reduction of the oxidation prod uct during the negative portion of the cyclic scan of E\. All detection process es pertinent to this article are irrevers ible; the oxidation products cannot be detected cathodically. However, even for a reversible redox system, there is sufficient loss of soluble oxidation products from the diffusion layer at the electrode by convective-diffusional mass transport so that the cathodic charge from reduction of detection products will not be equivalent to the anodic charge from the detection pro cess. Chromatographic applications Polyalcohols and carbohydrates (Mode I). All aldehydes, simple alco hols, glycols, polyalcohols, and carbo hydrates can be detected by pulsed
Several requirements pertaining to the cyclic sweep of E\ in IPAD must be satisfied to achieve maximum success for applications to LC. First, the cyclic scan of E\ must begin and end at a value for which the electrode is free of surface oxide; Ε < — 0 . 1 V vs. SCE for Au in 0.1 M NaOH (see Figure 1). Sec ond, the value of E\ should not extend into the region for cathodic detection of dissolved 0 2 ; Ε < - 0 . 1 V vs. SCE (see Figure 1) if O2 is present. Finally, the positive scan must not extend beyond the value for anodic solvent break down; ~ + 0 . 6 V vs. SCE (see Figure 1). From the residual i-E curve for Au in Figure 1, it is clear that only a small potential region centered at 0.1 V vs. SCE is appropriate to satisfy the first two criteria in 0.1 M NaOH con 592 A · ANALYTICAL CHEMISTRY, VOL. 62, NO. 10, MAY 15, 1990
electrochemical detection at Au and P t electrodes in alkaline media (pH > ~12). However, use of Au electrodes has the distinct advantage that detec tion can be achieved without simulta neous reduction of dissolved O2. The maximum anodic signal for polyalco hols and carbohydrates in 0.1 M NaOH is obtained at Ε = +0.1 - +0.2 V vs. SCE, and a value in this range is chosen for E\ in the PAD waveform (Figures 2a and 2b). The primary detection re actions are given by Equations 1-3 (see box) for glucose (40). The first step is a very fast two-elec tron oxidation of the aldehyde group to the corresponding carboxylic acid (Equation 1). This is followed by a se ries of relatively fast steps resulting in cleavage of the C1-C2 bond, with pro duction of HCO2H, followed by conver sion of the C2 and C6 to the correspond ing carboxylates (Equation 2). Addi tional sequential anodic cleavage of the remaining terminal carbons can occur according to Equation 3, but it occurs very slowly (40, 41). The observed number of electrons (rc0bs) for glucose detection at an Au electrode in a typi cal thin-layer LC detector is ~10 eq mol - 1 . At rates of convection higher than those that exist in the typical thin-layer LC detector, n0bs = ~ 8 eq mol - 1 . The limit of detection for glu cose by PAD (ii = 200 ms) is below 1 ng in 0.1 M NaOH with a linear response over more than 4 decades in concentra tion. The rates of anodic mechanisms at Au electrodes decline with increasing solution acidity, and virtually no re sponse for alcohols is obtained for pH « 12. The enhancement of oxidation
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Figure 4. Anion-exchange separation of carbohydrate mixture with gradient eiution. Detection: PAD (Mode A) at an Au electrode. E, = +0.1 V (i, = 610 ms, (, = 400 ms, t, = 200 ms); E2 = +0.8 V (fe = 120 ms); £3 = - 0 . 6 V (f3 = 300 ms). Column: Dionex AS-6 Carbopac. Solvents: A, 100 mM NaOH; B, 50 mM NaOH + 0.5 M NaOAc; C, H20. Eiution: isocratic (0-6 min) with 50% A + 50% C; linear (6-15 min) to 50% A + 50% B; isocratic (15-21 min) with 50% A + 50% B. Postcolumn addition: 0.4 M NaOH. Peaks: A, inositol; B, xylitol; C, sorbitol; D, mannitol; E, fucose; F, rhamnose; G, arabinose; H, glucose; I, xylose; J, fructose; K, sucrose; L, unknown; M, maltose; N, maltotriose; O, maltotetraose; P, maltopentaose; Q, maltohexaose; R, maltoheptaose.
rates by high alkalinity results from H + produced in reaction steps leading to the rate-determining step (40, 41). In contrast to the case of Au, useful analytical signals for alcohols and carbohydrates can be obtained at P t electrodes, even in concentrated acidic media ( 5 8). This is explained by the stronger adsorption of the intermediate reaction products at Pt, which has a lower electronic occupancy in surface d-orbitals than Au. The greatest convenience in LCPAD is realized when the separation phases can tolerate the conditions of pH and ionic strength desired for optimum detection. Polymeric phases are preferred because separation phases based on derivatized silica are not stable for extreme pH values. Anion-exchange phases have been applied successfully for separations of polyalcohols and carbohydrates (24, 25, 28,29). Carbohydrates are weakly acidic (42) and are adsorbed as anions from alkaline solutions (28). Hence, the order of eiution of carbohydrates correlates with pKa values, which decrease with increasing molecular weight. Retention
times for the anion-exchange separation of carbohydrates can be decreased conveniently by addition of acetate ion (OAc~). Olechno et al. (29) demonstrated the separation in 35 min of 43 components of a partially hydrolyzed dextrin sample using a linear solvent gradient from 0.1 M NaOH to 0.1 M NaOH + 0.6 M NaOAc. The first peak corresponded to glucose, and each succeeding peak represented a polyglucose molecule with a degree of polymerization increased by one (Peak 2 was for maltose [DP = 2], Peak 3 was for maltotriose [DP = 3], and so forth). Figure 4 illustrates the separation of a complex mixture of carbohydrates including sugar alcohols, monosaccharides, and several oligosaccharides. Sugar alcohols and monosaccharides were eluted under isocratic eiution with 50 mM NaOH (0-6 min). A linear solvent gradient was then applied (615 min) to bring the eluting phase to 75 mM NaOH + 250 mM NaOAc for eiution of xylose, fructose, sucrose, and maltose. Isocratic eiution continued (15-21 min) for the polyglucose series maltotriose (DP = 3) through malto-
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INSTRUMENTATION
Figure 5. Separation of a mixture of alcohols and glycols. Detection: PAD (Mode I) at a Pt electrode. Column: Dionex AS-1. Mobile phase: 0.050 M HCI04. Peaks: 1, glycerol; 2, ethylene glycol; 3, propylene glycol; 4, methanol; 5, ethanol; 6, 2-propanol; 7, 1-propanol; 8, 2-butanol; 9, 2-methyl-1-propanol; 10, 1-butanol; 11, 3-methyl-1-butanol; 12, 1-pentanol; 13, cyclohexanol; 14, 1-hexanol; 15, diethylene glycol.
heptaose (DP = 7). Larew et al. (43, 44) considered the challenge of quantitation of the chro matographic peaks for polyglucose compounds with DP = 2-7. A short (~1.5 cm) reactor column containing immobilized glucoamylase was insert ed into the LC system immediately af ter the separation column for conver sion of the polyglucose compounds to the equivalent amount of glucose. For 100% conversion of the components of each LC peak to the corresponding quantity of glucose, a single glucose calibration curve can suffice for peak area quantitation. The efficiency of conversion in the short reactor column was 87% for maltose (DP = 2), 93% for maltotriose (DP = 3), and 96% for the series DP = 4-7 (44). Undoubtedly, slightly longer reactor columns will produce more quantitative conversion for DP < 3. Application of the postseparation enzymatic reactor required addition of pH 5.5 buffer prior to the reactor to produce the optimum pH for enzymat ic activity. Furthermore, alkaline buff er was added following the reactor col umn to facilitate PAD at an Au elec trode. Subsequently, Larew and Johnson (45) demonstrated that for a solution having a low buffer capacity the high pH requirement of PAD can be temporarily achieved at Au elec trode surfaces as a result of the cathod-
ic dissolution of the surface oxide at E^ (i.e., AuO + H 2 0 + 2e" — Au + 20H") just prior to the detection process at E± in the next cycle of the waveform (45). Alcohols and glycols. Glycols are highly polar and thus are not amenable to GC. Furthermore, the absence of photometric detectability has rendered their sensitive detection very difficult in LC. The separation of a mixture of 15 alcohols and glycols in an anion-exchange column, using 0.050 M HCIO4 as the mobile phase, is shown in Figure 5 (46). Because of the acidic mobile phase, PAD was applied at a P t elec trode. The detection limit for ethylene glycol in water was determined to be - 1 0 ppb (S/N = 3). Alkanolamines. Alkanolamines are important in the chemical and pharma ceutical industries for production of emulsifying agents, corrosion inhibi tors, laundry additives, and dyes, and for purifying gases. These compounds lack natural chromophores and fluorophores for photometric and fluorometric detection; furthermore, their high polarity virtually eliminates use of GC for quantitative determinations. A reversed-phase separation was de veloped using a mixed acetonitrile-water mobile phase with sodium dodecanesulfonate (SDS) as an ion-pairing agent (47). Acetonitrile is adsorbed at noblemetal surfaces and therefore interferes dramatically with the amperometric
594 A · ANALYTICAL CHEMISTRY, VOL. 62, NO. 10, MAY 15, 1990
response for alcohols. However, virtu ally no interference from acetonitrile was observed for detection of the alco hol moiety of alkanolamines. An oxidecatalyzed amine signal observed for al kanolamines was proof for the adsorp tion of the amine even in the presence of acetonitrile. It is proposed that ano dic detection of the alcohol group in the presence of acetonitrile is a beneficial consequence of the amine adsorption that brings the associated alcohol group close to the electrode surface. This effect was observed even for 6-amine-l-hexanol with a 5-carbon chain separating the alcohol and amine groups. The interference of acetonitrile added postcolumn to the effluent stream during separations of alcohols (and polyalcohols and carbohydrates) can be applied to distinguish between peaks for alkanolamines and alcohols in complex mixtures. Chromatograms are shown in Figure 6 with detection of the alcohol group by Mode I for mixtures of linear, branched, and complex alkanolamines separated by reversed-phase ion-pair ing chromatography (47). Under the conditions used, monoethanolamine and triethanolamine were well re-
Figure 6. Ion-pairing, reversed-phase separation of alkanolamines. Detection: PAD (Mode I) at an Au electrode. Mo bile phase: 20% acetonitrile with 2 mM SDS. Postcolumn addition: 0.2 M NaOH. Column: Wa ters C-18 μΒοηάβρβ^ Peaks: A, 2-amino-1ethanol; B, 3-amlno-1-propanol; C, 4-amlno-1-butanol; D, 5-amlno-1-pentanol; E, 6-amlno-1-hexanol; F, 2-amlno-1-propanol; G, 2-amlno-1butanol; H, 2-amino-1-pentanol; I, 1-amino-2propanol; J, 2-amlno-2-methyl-1-propanol; K, 2-amino-1-phenylethanol. (Adapted from Ref erence 47.)
Figure 7. An ion-exchange separation of sugars and amino sugars. Detection: PAD (Mode I) at an Au electrode. Col umn: Dionex HPIC-AS6A. Mobile phase: 0.2 mM NaOH. Postcolumn addition: 0.2 M NaOH. Peaks: A, fucose; B, 2-deoxyribose; C, galactosamine; D, galactose; E, glucosamine; F, glucose; G, mannose; H, Λί-acetylglucosamine; I, W-acetylgalactosamine.
solved; however, monoethanolamine and diethanolamine were co-eluted. Postcolumn addition of 0.2 M NaOH was used to ensure the high pH needed for optimum sensitivity for PAD at an Au electrode. The detection limit (S/N = 3) for ethanolamine was deter mined to be 40 ppb (8 ng in the 200-μΙ_. sample injected) with linear response from 40 ppb to ~10 ppm. Sugar amines. Anodic detection of sugar amines can be based on Mode I (the carbohydrate) or Mode II (the ad sorbed amine). Mode I yields greater detectability. The quantitative separa tion and detection of sugar amines are important in the pharmaceutical in dustry, especially in antibiotic produc tion (48) and in glycoprotein research within biological disciplines (49-51). LC-PAD (Mode I) results are shown in Figure 7 to illustrate the anion-exchange separation of a mixture of nine compounds, eight of which are typical ly found in hydrolyzates of glycopro teins; 2-deoxyribose was added as an internal standard (52). An interesting effect results from the postcolumn addition of acetonitrile. Peaks for simple sugars (e.g., glucose) are extensively attenuated. This same effect occurs for sugar amines in which the amine groups are acetylated (e.g., iV-acetylglucosamine). Peaks for sugar amines are not attenuated by acetoni trile when the amine groups are sterically free to adsorb at the electrode sur face. This phenomenon is considered to have the same origins as in the case of the acetonitrile effect on alcohol and alkanolamine peaks (e.g., the strong adsorption of the amine groups pre vents interference by adsorbed aceto nitrile in the surface-catalyzed detec-
Figure 8. Isocratic separation of three amino acids. Electrode: Au with SCE reference. Column: Dionex HPIC-AS6. Mobile phase: 0.10 M NaOH. Detection: (a) PAD; (b) and (c) IPAD. Peaks: A, lysine; B, asparagine; C, 4-hydroxyproline. Concentrations: (a) and (b) ~ 5 0 ng each; (c) ~0.5 ng each (i.e., ~ 3 pmol).
tion mechanism). Amino acids. Most amino acids in biological materials are aliphatic and have been perceived as not electroactive (53, 54). Their detection in LC has commonly been achieved by photomet ric absorbance after derivatization using ninhydrin reagent, by fluores cence using o-phthaldehyde reagent, or by amperometric detection of the phenyl- or methylthiohydantoin adducts at Ag and Hg electrodes. Amino acids can be detected directly at P t (26) and Au (33) electrodes in alkaline solutions by the oxide-cata lyzed mechanism of Mode II. The Au electrode is preferred over P t to mini mize interference from dissolved O2. Figures 8a and 8b show a comparison of PAD and IPAD, respectively, at an Au electrode for the isocratic separation of three amino acids on an anion-exchange column (46). Clearly, IPAD is
preferred over PAD to minimize base line offset and drift. Figure 8c shows the LC-IPAD results for the same three compounds at their detection limits (0.5 ng, ~ 3 pmol). It was not possible to find isocratic conditions for resolution of all amino acids. A gradient procedure was devel oped that incorporated a change in pH (33, 55). A pH change causes a shift in the onset of the anodic wave for oxide formation (Wave A in Figure 1) by the amount ~—60 mV p H - 1 . Consequent ly, the baseline obtained for PAD in creases (anodically) for an increase in pH. The use of IPAD results in a signif icant decrease in the baseline shift for small changes in pH (i.e., pH < 2). However, optimum potential values for the IPAD waveform also shift with change in pH ( 60 mV pH" 1 ). The negative consequences of this fact can be decreased significantly by use of a
ANALYTICAL CHEMISTRY, VOL. 62, NO. 10, MAY 15, 1990 · 595 A
INSTRUMENTATION glass membrane, H + -selective elec trode, as the reference in place of the conventional pH-independent elec trodes (e.g., Hg/Hg 2 Cl 2 or Ag/AgCl ref erence electrodes [56,57]). Because the pH dependence of the glass electrode is the same as for the processes at the Au electrode, the voltammetric response at the Au electrode appears to be pH independent. Chromatographic results are shown in Figure 9 for the gradient separation of 20 amino acids present in a protein hydrolyzate. Detection was by IP AD at an Au electrode with a glass membrane reference electrode {33). Detection lim its (S/N = 3) by IP AD were determined to be ~ 3 - 5 pmol (46). It is especially significant that the sensitivity of IP AD is approximately the same for primary and secondary amino acids. Sulfur compounds. Numerous or ganic and inorganic sulfur compounds are adsorbed at the oxide-free surfaces of Au and P t electrodes and can be de tected by Mode II (34-37). They in clude thioalcohols, thioethers, thiophenes, thiocarbamates, organic thiophosphates, and numerous inorganic compounds. Adsorption is prerequisite to detection; therefore, at least one nonbonded electron pair must reside on the sulfur atom. Hence, sulfonic ac ids and sulfones are not detected. LCPAD has been demonstrated for a mix-
ture of sulfur-containing pesticides separated by reversed-phase LC (37) with a mobile phase composed of 50% acetonitrile in aqueous acetate buffer (pH 5). The kinetics for detection of adsorbed sulfur compounds are quite favorable at pH 5, and it was not neces sary to rely on postcolumn addition of NaOH. Conclusion
The analytical significance of PAD and IPAD at Au and P t electrodes is the sensitive detection of numerous ali phatic compounds separated by LC. As illustrated, PAD and IPAD are com patible with several forms of LC, in cluding ion-exchange, ion-pairing, ionexclusion, and reversed-phase chroma tography. We speculate t h a t only application to normal-phase LC will be problematic because postcolumn addi tion of the aqueous buffer solution re quired to support the detection pro cesses could result in serious loss of sol ubility of various analytes to be separated. We anticipate that significant devel opments in PAD and IPAD will occur for detection of yet-to-be explored compounds, including aromatic com pounds. Current research in our lab oratory also is being directed to the de tection of macromolecules, including peptides and small proteins, and for applications of PAD and IPAD at microcylindrical (wire) electrodes for use in capillary-bore LC columns. References
Figure 9. Ion-exchange separation of amino acids in hydrolyzed protein. Detection: IPAD (Mode II) at an Au electrode. Ref erence: glass membrane pH electrode. Column: Dionex AS-8 with AG-6 guard column. Gradient elution: described in Reference 33. Peaks: 1, arginine; 2, lysine; 3, glutamine; 4, asparagine; 5, threonine; 6, alanine; 7, glycine; 8, serine; 9, va line; 10, proline; 11, isoleucine; 12, leucine; 13, methionine; 14, histidine; 15, phenylalanine; 16, glutamic acid; 17, aspartic acid; 18, cysteine; 19, cystine; 20, tyrosine. (Adapted from Reference 33.)
(1) Chemical Deriuatization in Analytical Chemistry: Chromatography; Frei, R. W.; Lawrence, J. F., Eds.; Plenum Press: New York, 1981; Vol. 1. (2) Post-Column Reaction Detectors in HPLC; Krull, I. S., Ed.; Marcel Dekker: New York, 1986. (3) Adams, R. N. Electrochemistry at Solid Electrodes; Marcel Dekker: New York, 1969. (4) Shoup, R. E. Recent Reports on Liquid Chromatography-Electrochemistry; Bioanalytical Systems Press: West Lafayette, IN, 1982. (5) Oilman, S. J. Phys. Chem. 1963, 67, 7884. (6) Breiter, M. W. Electrochim. Acta 1963, 8, 973-83. (7) Giner, J. Electrochim. Acta 1964, 9, 6377. (8) Oilman, S. In Electroanalytical Chem istry; Bard, A. J., Ed.; Marcel Dekker: New York, 1967; Vol. 2, pp. 111-92. (9) Woods, R. In Electroanalytical Chem istry; Bard, A. J., Ed.; Marcel Dekker: New York, 1976; Vol. 9, pp. 20-27. (10) Hammett, L. P. J. Am. Chem. Soc. 1924,46, 7-19. (11) Armstrong, G.; Himsworth, F. R.; But ler, J.A.V. Proc. R. Soc. London A 1934, 143, 89-103. (12) Clark, D.; Fleishman, M.; Pletcher, D. J. Electroanal. Chem. 1972, 36,137-46. (13) MacDonald, Α.; Duke, P. D. J. Chromatogr. 1973,83, 331-42.
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(14) Stulik, W.; Hora, V. J. Electroanal. Chem. 1976, 70, 253-63. (15) Fleet, B.; Little, C. J. J. Chromatogr. Sci. 1974,12, 747-52. (16) van Rooijen, H. W.; Poppe, H. Anal. Chim. Acta 1981,130, 9-22. (17) Ewing, A. G.; Dayton, Μ. Α.; Wightman, R. M. Anal. Chem. 1981, 53, 184247. (18) Berger, T. A. U.S. Patent 4 496 454, Jan. 29, 1985. (19) Tenygl, J. In Electrochemical Detec tors; Ryan, T. H., Ed.; Plenum Press: New York and London, 1984; pp. 89-103. (20) Hughes, S.; Meschi, P. L.; Johnson, D. C. Anal. Chim. Acta 1981,132, 1-10. (21) Hughes, S.; Johnson, D. C. Anal. Chim. Acta 1981,132,11-22. (22) Hughes, S.; Johnson, D. C. J. Agric. Food Chem. 1982, 30, 712-14. (23) Hughes, S.; Johnson, D. C. Anal. Chim. Acta 1983,149,1-10. (24) Rocklin, R. D.; Pohl, C. A. J. Liq. Chromatogr. 1983, 6, 1577-90. (25) Edwards, P.; Haak, Κ. Κ. Amer. Lab. 1983, April, 78-87. (26) Johnson, D. C. Nature 1986, 321, 45152. (27) Neuburger, G. G.; Johnson, D. C. Anal. Chem. 1987, 59, 203-04. (28) Edwards, W. T.; Pohl, C. Α.; Rubin, R. Tappi J. 1987, 70,138-40. (29) Olechno, J.; Carter, S. R.; Edwards, W. T.; Gillen, D. G. Amer. Biotech. Lab. 1987 5 38-50. (30) Austin, D. S.; Polta, J. Α.; Polta, T. Z.; Tang, A.P-C; Cabelka, T. D.; Johnson, D. C. J. Electroanal. Chem. 1984, 108, 227-48. (31) Polta, J. Α.; Johnson, D. C. J. Liq. Chromatogr. 1983, 6, 1727-43. (32) Johnson, D. C; Polta, J. Α.; Polta, T. Z.; Neuburger, G. G.; Johnson, J.; Tang, A.P-C; Yeo, I-H.; Baur, J. J. Chem. Soc, Faraday Trans. 1 1986,82,1081-98. (33) Welch, L. E.; LaCourse, W. R.; Mead, D. Α., Jr.; Johnson, D. C. Anal. Chem. 1989,61, 555-59. (34) Polta, T. Z.; Johnson, D. C. J. Elec troanal. Chem. 1986, 209,159-69. (35) Polta, T. Z.; Johnson, D. C; Luecke, G. R. J. Electroanal. Chem. 1986, 209, 171-84. (36) Johnson, D. C; Polta, T. Z. Chroma togr. Forum 1986,1, 37-43. (37) Ngoviwatchai, Α.; Johnson, D. C. Anal. Chim. Acta 1988,275,1-12. (38) Neuburger, G. G.; Johnson, D. C. Anal. Chim. Acta 1987,192, 205-13. (39) Neuburger, G. G.; Johnson, D. C. Anal. Chem. 1988,60, 2288-93. (40) Larew, L. Α.; Johnson, D. C. J. Elec troanal. Chem. 1989,262, 167-82. (41) Larew, L. Α.; Johnson, D. C, unpub lished data, Iowa State University, Ames, IA, Nov. 1989. (42) Rendleman, J. A. In Advances in Chemistry Series §117; Gould, R. F., Ed.; American Chemical Society: Washington, DC, 1973; p. 51. (43) Larew, L. Α.; Mead, D. Α., Jr.; John son, D. C. Anal. Chim. Acta 1988,204,4351. (44) Larew, L. Α.; Johnson, D. C. Anal. Chem. 1988, 60, 1867-72. (45) Larew, L. Α.; Johnson, D. C. J. Elec troanal. Chem. 1989,264,131-47. (46) LaCourse, W. R.; Johnson, D. C, un published data, Iowa State University, Ames, IA, Nov. 1989. (47) LaCourse, W. R.; Jackson, W. Α.; Johnson, D. C. Anal. Chem. 1989, 61, 2466-71. (48) Polta, J. Α.; Johnson, D. C; Merkel, K. E. J. Chromatogr. 1985, 324, 407-14. (49) Hardy, M. R.; Townsend. R. R. Proc.
Nat. Acad. Sci. 1988,85, 3289-93. (50) Hardy, M. R.; Townsend, R. R.; Lee, Y. C. Anal. Biochem. 1988,170, 54-62. (51) Townsend, R. R.; Hardy, M. R.; Hindsgaul, O.; Lee, Y. C. Anal. Biochem. 1988,274,459-70. (52) Jackson, W. Α.; LaCourse, W. R.; Johnson, D. C, Iowa State University, Ames, IA, unpublished results. (53) Malfoy, M.; Reynaud, J. A. J. Electroanal. Chem. 1980,114, 213-23. (54) Joseph, H. M.; Davies, P. Curr. Sep. 1982,4,62-65. (55) Slingsby, R. Α.; Dionex Corp., Sunny vale, CA, personal communication, 1989. (56) Mead, D. Α.; Larew, L. Α.; LaCourse, W. R.; Johnson, D. C. In Advances in Ion Chromatography; Jandik, P.; Cassidy, R. M., Eds.; Century International: Franklin, MA, 1989; Vol. 1, pp. 13-34. (57) LaCourse, W. R.; Mead, D. Α., Jr.; Johnson, D. C. Anal. Chem. 1990,62,22024.
Recommended reading Adams, R. N. Electrochemistry at Solid Electrodes; Marcel Dekker: New York, 1969. Laboratory Techniques in Electroanalytical Chemistry; Kissinger, P. T.; Heineman, W. R., Eds.; Marcel Dekker: New York, 1984. Weber, S. G. In Detectors for Liquid Chro matography; Yeung, E. S., Ed.; John Wi ley and Sons: New York, 1986; pp. 229-91.
Johnson, D. C; Weber, S. G.; Bond, A. M.; Wightman, R. M.; Shoup, R. E.; Krull, I. S. Anal. Chim. Acta 1986,180,187-250. Stulik, K.; Pacakova, V. Electroanalytical Measurements in Flowing Solutions; John Wiley and Sons: New York, 1987. Austin-Harrison, D. S.; Johnson, D. C. Electroanalysis 1989,1,189-97.
Dennis C. Johnson received a B.A. de gree in chemistry from Bethel College (St. Paul, MN) in 1963 and a Ph.D. in analytical chemistry from the Univer sity of Minnesota in 1967. He joined the faculty at Iowa State University in 1968. His research interests focus on the electrocatalysis of anodic reac tions at noble-metal electrodes and at pure and mixed-metal oxide elec trodes for applications with chromato
graphic detection, electrochemical synthesis, and the anodic degradation of toxic waste chemicals ("electro chemical incineration").
William R. LaCourse received an A.S. degree in chemical engineering from Thames Valley State Technical Col lege (Norwich, CT) in 1977, a B.S. de gree in chemistry from Charter Oak College (Hartford, CT) in 1982, and a Ph.D. in analytical chemistry from Northeastern University in 1987. He has worked in the pharmaceutical in dustry on the development of product assays and with Johnson's LC-PAD program since 1987. His interests in clude basic and applied research on electrochemical detection in LC.
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