Uptake, Speciation, and Uncoupling Activity of Substituted Phenols in

phorylation or photophosphorylation. In this study, the relationship between uncoupling activity, total concentration, and speciation in the photosynt...
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Environ. Sci. Technol. 1996, 30, 3071-3079

Uptake, Speciation, and Uncoupling Activity of Substituted Phenols in Energy Transducing Membranes BEATE I. ESCHER, MARIO SNOZZI, AND RENE ´ P. SCHWARZENBACH* Swiss Federal Institute for Environmental Science and Technology (EAWAG), and Swiss Federal Institute of Technology (ETH), CH-8600 Du ¨ bendorf, Switzerland

Phenolic compounds are toxic to many organisms in that they may affect the energy production in cells by inhibition of the electron transport or by destroying the electrochemical proton gradient built up across membranes. This latter mode of toxic action is commonly referred to as uncoupling of oxidative phosphorylation or photophosphorylation. In this study, the relationship between uncoupling activity, total concentration, and speciation in the photosynthetic membrane (chromatophores) of the purple bacterium Rhodobacter sphaeroides has been evaluated for 18 nitro- and chlorophenols covering a wide range of hydrophobicity and acidity. The uncoupling activity has been determined by time-resolved spectroscopy and is quantified by a pseudo-first-order rate constant, kobs, which is a measure for the increased decay rate of the membrane potential in the presence of a certain amount of a given phenol. The experimental data can be described by an extended “shuttle mechanism” model in which it is assumed that the rate of diffusion of the phenoxide and/or a phenoxide/phenol-heterodimer species through the lipid bilayer of the membrane determines the rate of decay of the electrochemical proton gradient: kobs AAHA AHA , where C cph and C cph are ) k1C cph + k 2′ C cph C cph the concentrations of the phenoxide and phenol, respectively, in the chromatophores (both estimated from membrane-water partitioning experiments), k1 is a measure of the mobility of the phenoxide in the lipid bilayer; and k2′ is a lumped parameter describing both the tendency of the compound to form a heterodimer in the membrane as well as the mobility of this heterodimer in the lipid bilayer. To our knowledge, this is the first study in which, for a given class of ionogenic organic compounds, a direct quantitative measure of a specific toxic effect (i.e., uncoupling) has been successfully related to the actual concentration and speciation of the compounds at the target site (i.e., in the membrane). This study

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 1996 American Chemical Society

demonstrates that it is possible to separate the contributions of uptake, speciation, and actual activity (expressed by k1 and/or k2) to the overall uncoupling potency of a given phenol, which is necessary for the derivation of improved quantitative structure-activity relationships (QSARs). Furthermore, the approach taken in this study offers the possibility to evaluate quantitatively synergistic and antagonistic effects of different phenolic compounds on energy transduction when such compounds are present in mixtures.

Introduction Phenolic compounds are ubiquitous contaminants that are of considerable environmental concern because they are toxic to many organisms by interfering with one of the basic cell functions, namely, energy transduction. In energytransducing membranes, substituted phenols may act as uncouplers in that they can destroy the electrochemical proton gradient by transporting protons back across the membrane (1, 2), and/or they may inhibit the electron flow by binding directly to specific components of the electron transfer chain (3). In addition, by simply accumulating in the membrane, less active phenols can nonspecifically perturb the membrane functions, thus causing a so-called narcotic effect. In a parallel paper (4), we have presented a new method for the determination of the uncoupling and inhibitory activities of organic compounds in energy-transducing membranes. This method has the advantage that both processes, uncoupling and inhibition, can be quantified independently with the same experimental setup. Furthermore, the method is relatively simple and fast. Chromatophores of the photosynthetic bacterium Rhodobacter sphaeroides were chosen as a model system. The membranes of these vesicles contain a simple cyclic electron transfer chain (5). Short flashes of light induce photooxidation of the reaction center followed by electron and proton transfer leading to a buildup of an electrochemical proton gradient. By measuring the membrane potential as a function of time after a single-turnover flash, the uncoupling process can be detected and quantified. The membrane potential is the major component of the electrochemical proton gradient in chromatophores and is detected via an electrochromic shift of the absorption band of carotenoids in the light-harvesting antennae (6). Protonophoric uncouplers increase the decay rate of the membrane potential. This decay rate can be described by first-order kinetics, thus the uncoupling activity can be expressed by a pseudo-first-order rate constant. For the phenols investigated, the aqueous concentrations required to observe significant uncoupling were in the micromolar range. By following spectroscopically the redox kinetics of several components of the electron transfer chain, namely, reaction center, as well as b- and c-type cytochromes, it was also possible to localize the site of inhibition and to * Corresponding author e-mail address: [email protected]; telephone 41-1-823 5109; fax 41-1-823 5471.

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quantify the inhibitory effect (4). It was found that phenolic compounds compete with molecules from the quinone pool for the quinone reductase site Qi of the cytochrome bc1 (cyt bc1) complex. The binding was reversible, thus the turnover of the cyt bc1 complex was only slowed down but not totally blocked. The aqueous concentrations of phenols required for 50% of maximum inhibition were however on the order of 10-4 M and higher, so that in most cases inhibition was less important than uncoupling. The concentrations indicated above refer to the total concentrations present in the aqueous phase. However, for relating the activity of a given compound to its structure or properties, it is crucial to know what fraction of the compound actually reaches the target site(s), in this case, the energy-transducing membrane. An estimate of the concentration at the target site is even more important for hydrophobic ionogenic compounds because neutral and charged species are not taken up to the same extent. In the protonophoric shuttle mechanism of uncoupling (1), both species have to act together to cause effective proton transport. There are also indications that the phenoxide is the main active species for inhibition (3, 7). Consequently, the speciation inside the membrane is a pivotal factor for the toxic potency of phenolic compounds and, therefore, needs further investigation. Until recently, it was commonly assumed that the dissociated form of phenolic compounds, the phenoxide, plays a minor role in the overall uptake (8) or in the toxic effect (9, 10). This assumption was mainly based on the fact that the octanol-water distribution ratio of the negatively charged phenoxide is about 3 orders of magnitude smaller than the Kow of the corresponding neutral species (11, 12). However, in previous work, we have shown that partitioning into the bulk solvent octanol significantly underestimates the extent of phenoxide partitioning between water and biological membranes (13). For a series of chlorinated and nitrated phenols, the differences between the membrane-water distribution ratios of the neutral and the corresponding anionic species were less than 1 order of magnitude. Furthermore, we have demonstrated that liposome-water distribution ratios, Dlipw, can be used to assess the total concentrations of phenols in energytransducing membranes and that the speciation of phenols inside the lipid bilayer can be estimated by evaluation of the pH dependence of Dlipw (13). In this paper, we use the results of liposome-water partitioning experiments to evaluate the relationship between uncoupling activity, total concentration, and speciation of a series of phenolic compounds in the membranes of the purple photosynthetic bacterium Rb. sphaeroides. To this end, a mechanistic two-parameter model is developed for describing the observed uncoupling activities. The model is based on the assumption that the rate of transport of the phenoxide and/or a phenoxide/ phenol heterodimer species through the lipid layer of the membrane determines the rate of decay of the electrochemical proton gradient. The model is validated by experimental data and is then used to interpret the uncoupling activities determined by time-resolved spectroscopy for 18 nitro- and chlorophenols covering a wide range of hydrophobicity and acidity. The results of this study provide important insights into the molecular factors that determine the uncoupling potency of phenols and form an important base for the development of more meaningful structure-activity relationships for such compounds.

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FIGURE 1. Mechanistic model of the protonophoric activity of phenolic compounds in chromatophore membranes. Photooxidation followed by electron transfer causes the buildup of ∆µH+, which is equivalent to ∆ψ in chromatophores (25). Equilibrium processes: partitioning of HA and A- into the lipid bilayer described by Kcphw(HA) and Dcphw(A-), formation of the heterodimer, KAHA-. Diffusion processes: diffusion of HA, JHA, of A-, JA-, and of AHA-, JAHA-.

Model for Describing the Protonophoric Activity of Phenols in Chromatophores In this section, a simple model is developed for describing the processes that are likely to occur in the chromatophores of Rb. sphaeroides after a single turnover flash in the presence of phenolic compounds or other weak organic acids. The conceptual framework is based on the “shuttle mechanism” approach proposed by McLaughlin and coworkers (14, 15), who evaluated the protonophoric activity of phenylhydrazones in artificial planar lipid bilayers. The model presented here differs from that of McLaughlin and co-workers in that (i) not only one but two charged species inside the membrane are considered, [i.e., the phenoxide, Acph , and a heterodimer, AHAcph , which may be formed by association (i.e., hydrogen bonding) of a phenoxide and a phenol species inside the membrane (16)] and (ii) the actual concentrations of these species in the chromatophores are considered. Figure 1 summarizes the various species and processes taken into account. The model presumes that, at equilibrium, which is described by the equilibrium distribution ratios Kcphw(HA) and Dcphw(A-), all sorbed phenolic species are located near the hydrophobic-hydrophilic interface in the membrane (17). After the flash-induced photooxidation of the reaction center in the chromatophore, the electrochemical proton gradient, ∆µH+, which is built up by coupled electron and proton transfer, is diminished by the following sequence of processes: (1) Driven by the potential gradient, ∆ψ, Acph and/or AH Acph diffuse through the membrane from the n-side to the p-side thereby decreasing ∆ψ. (2) At the p-side, Acph and/or AHAcph pick up a proton from the adjacent aqueous boundary layer thus forming neutral HAcph. (3) Since processes 1 and 2 form a concentration gradient across the membrane, an equivalent number of HAcph

diffuse to the n-side and release a proton to the aqueous boundary layer. This completes the “shuttle cycle” that leads to the net transfer of a proton across the membrane. As discussed in detail elsewhere (4, 7), the decay of ∆ψ (and hence ∆µH+) can be viewed as the discharge of an electrical capacitor through a constant resistance and can thus be quantified by a pseudo-first-order rate constant, kobs. If one now assumes that, among all the processes summarized in Figure 1, the diffusion of Acph and AHAcph through the lipid bilayer primarily determines the decay rate (14), than in the most simple approach, kobs can be expressed by the linear combination of two terms, each describing the contribution of the corresponding species:

kobs ) k1[A-]cph + k2[AHA-]cph

(1)

where [A-]cph and [AHA-]cph are the equilibrium concentrations of the phenoxide and the heterodimer, respectively, in the chromatophore. By defining an equilibrium constant, KAHA-, for the formation of the heterodimer in the chromatophore membrane:

KAHA- )

[AHA-]cph [HA]cph[A-]cph

(2)

eq 1 can be rewritten as

kobs ) k1[A-]cph + k′2[HA]cph[A-]cph

(3)

where k′2 ) k2KAHA-. In terms of total phenol concentration in the chromatophore, Ccph ) [HA]cph + [A-]cph + 2[AHA-]cph, and by assuming that [AHA-]cph , [HA]cph, [A-]cph, eq 3 can also be expressed as cph cph 2 kobs ) k1f Acph - Ccph + k′2f A- (1 - f A- )C cph

(4)

where f Acph is the fraction of total phenol inside the membrane that is present as phenoxide, and 1 - f Acph - () cph f HA ) is the fraction present as non-dissociated phenol. Since Ccph and f Acph - can be estimated from liposome-water partition ratios (see Materials and Methods), k1 and k′2 can be derived from the experimental data by a second-order polynomial fit of kobs versus Ccph: 2 kobs ) aCcph + bC cph

(5)

cph cph where a ) k1f Acph - and b ) k′2f A- (1 - f A- ). In this model, k1 is interpreted as a measure of the mobility of the phenoxide in the lipid bilayer, while k′2 is a composite measure of the mobility of AHA- and of the tendency of the phenol and phenoxide to form a heterodimer in the chromatophore membrane. Note that with the experimental data available, the two factors that determine k′2 cannot be quantified separately.

Materials and Methods Chemicals. The phenols (full names and abbreviations are given in Table 1) were purchased from the following companies: Riedel-de Ha¨en (Seelze, Germany): 24DCP, 26DCP, 245TriCP, 246TriCP, 2346TeCP, DNOC, Dinoseb, and Dino2terb; Fluka (Buchs, Switzerland): 345TriCP, 2345TeCP, PCP, 4NP, 24DNP, and 34DNP; Aldrich (Buchs,

Switzerland): 26DNP, DNPC, and Dino4terb; Merck (Zu ¨rich, Switzerland): 34DCP. The following biological buffers were used: MES [2-(N-morpholino)ethanesulfonic acid, pKa ) 6.15]; MOPS [3-(N-morpholino)propanesulfonic acid, pKa ) 7.2]; HEPPS [N-(2-hydroxyethyl)piperazine-N′-3-propanesulfonic acid, pKa ) 7.8]; CHES [2-(cyclohexylamino)ethanesulfonic acid, pKa ) 9.55], all of which were from Fluka (Buchs, Switzerland). Materials used for kinetic spectroscopy are described in refs 4 and 7. Preparation and Characterization of the Chromatophores. Chromatophores of the purple bacterium Rodobacter sphaeroides were prepared as decribed previously (4, 7). The concentration of the components of the reaction center complex was determined with the method of timeresolved spectrophotometry by measuring the difference between the extinction coefficients of the oxidized and reduced species at 542 nm. At this wavelength, the extinction coefficient ∆542nm of the reduced minus oxidized species is 10300 M-1 cm-1 (18). The phospholipid content, [PL]cph, of the chromatophore suspension was determined spectrophotometrically at 623 nm after mineralization to inorganic phosphate and formation of a malachite green/ molybdenum/phosphate complex according to Petitou (19). Determination of the Uncoupling Activity. Timeresolved spectroscopy was used to determine the uncoupling activity of phenolic compounds. The kinetic singlebeam spectrophotometer equipped with a flash excitation unit is described in detail elsewhere (4, 7). The measurements were performed in an anaerobic cuvette at a redox potential of 120-130 mV, held constant by poising with sodium dithionite and ferricyanide. The pH was adjusted using MES, MOPS, HEPPS, or CHES buffer or a mixture thereof. The total buffer and K+ concentrations (KCl/KOH) were 50 and 100 mM, respectively. For the uncoupling experiments, the stock suspension of chromatophores (pH 7.0) was diluted in an appropriate buffer, redox mediators were added, and the suspension was equilibrated in the dark for at least 1 h (or 30 min for pH 7.0). During this time, the redox potential was poised to 120-130 mV with sodium dithionite and ferrycyanide. Then, several measurement cycles with increasing concentrations of phenol were performed. During one measurement cycle, 4-8 kinetic traces were averaged, each of which consisted of the relative absorption at 503 nm 50 ms before and 1 s after a short (2-µs) xenon flash, followed by at least 1 min of re-equilibration. The concentration of the chromatophore suspension was adjusted so that the xenon flash induced single turnover of approximately 95% of the reaction centers. The detection wavelength of 503 nm was chosen because the absorbance change due to the electrochromic band shift of the carotenoids (6) is large and not disturbed by other signals. The absorbance change at 503 nm was shown to be approximately proportional to the membrane potential (20). Figure 2 shows an example of the original kinetic traces in the absence (i.e., the control) and in the presence of a certain amount of a phenol (e.g., Dinoseb). From these data, kobs can be derived by a linear regression of ln (x(t)) versus time (see insert in Figure 2):

ln(x(t)) ) -kobst

(6)

where x(t) ) ∆A503,phenol/∆A503,control(t). Note that this procedure yields a value of kobs that is effectively nor-

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ments. With this assumption, the liposome-water distribution ratios of the neutral and anionic species, respectively, are defined as

Klipw(HA) )

[HA]lip [HA]w

(9)

and

Dlipw(A-) )

FIGURE 2. Original kinetic traces of the control (i.e., without uncoupler added) and with 125 µmol kgcph-1 Dinoseb added. Insert: The slope of ln x(t) versus time corresponds to the observed rate constant of uncoupling kobs. [x(t) ) ∆A503(Dinoseb)/∆A503(control)].

malized (via the control) for the properties of a particular chromatophore preparation. For more details see ref 4. Estimation of the Total Concentration and the Speciation of the Phenols in the Chromatophores. In a previous study (13), we have shown that liposomes constructed from zwitterionic phosphatidylcholine are appropriate model systems for mimicking the partitioning behavior of both phenol and phenoxide species between the chromatophores of Rb. sphaeroides and water. Liposomes were chosen as model systems because they are much easier to prepare than chromatophores, and even more importantly, they are much more robust with respect to changes in solution conditions, in particular to changes in pH. At a given pH and salt concentration (see below), the liposome-water distribution ratio, Dlipw, which is defined as the ratio of the sum of the concentrations of all phenol species in the lipid phase (i.e., neutral phenol, phenoxide, and possibly the ion pair, in this case potassium phenoxide) divided by the total phenol concentrated in the aqueous phase, can be expressed by (4):

Dlipw(pH) )

Clip w ) f HA Klipw(HA) + f Aw-Dlipw(A-) (7) Cw

where Clip ) [HA]lip + [A-]lip + [AK]lip, Cw ) [HA]w + [A-]w, w and f HA and f Aw- are the fractions of the phenol in the nondissociated and dissociated form, respectively, in the aqueous phase: w f HA )

1 1 + 10

(pH-pKaw)

w f Aw- ) 1 - f HA

(8)

and Kaw is the acidity constant. Note that for the interpretation of the uncoupling effects, (or Acph , see no distinction is made between the anion Alip below) and a possible ion pair AKlip (or AKcph), because it is not clear to what extent the ion pair really exists in the membrane and because it can be assumed that the ion pairing is unimportant at the ionic strength of the experi

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[A-]lip [A-]w

(10)

where [A-]lip encompasses both A- and AK. Klipw (HA) and Dlipw(A-) can be derived from determinations of Dlipw at various pH values (13). Note that Dlipw(A-) is dependent on the K+ concentration. Therefore, all uncoupling experiments were conducted at the same K+ concentration (100 mM) as was used for the liposome-water partitioning experiments. For estimating the chromatophore-water distribution ratios from the liposome-water distribution ratios, it can now be assumed that the phenolic species partition predominantly into the lipid bilayer part of the chromatophores (13). Thus, the chromatophore-water distribution ratios can be obtained by simply multiplying the corresponding liposome-water distribution ratios (eq 7) with the fraction of lipids present in the chromatophores, which in this case was 0.3:

Dcphw(pH) )

Ccph ) 0.3Dlipw(pH) Cw

(11)

Table 1 summarizes the Dcphw values of the model compounds for a solution pH of 7.0. Hence, at a given pH, Ccph is calculated from the measured total concentration in the aqueous phase by

Ccph ) Dcphw(pH)Cw

(12)

For estimating the fraction of phenoxide species, f Acph - , present in the chromatophores at a given aqueous pH (see eq 4), an operational acidity constant, Kacph, in the chromatophore phase is defined. This operational acidity constant can be viewed as the pH of the adjacent aqueous phase at which the ratio of phenol and phenoxide in the chromatophore equals to one. Hence, Kacph is defined as (see also ref 7):

[A-]cph Dcphw(A-) ) Kaw Kacph ) [H+]w [HA]cph Kcphw(HA)

(13)

For a given pH, the fraction of the total phenol present as anionic species in the chromatophore can then be estimated by

f Acph - ) 1+

1 1 ) [HA]cph 1 + 10(pKacph - pH)

(14)

[A-]cph

The Kacph values as well as the f Acph - values for pH ) 7.0 of compounds investigated are also given in Table 1.

TABLE 1

Properties Relevant for Assessing the Uptake and Speciation of Substituted Phenols in Chromatophore Membranes at pH 7.0a compound

abbreviation

pKaw b

pKacph c

-1))d log(Dcphw/(L kgcph

cph e f A-

cph f f A(1 - f Acph - )

2,4-dichlorophenol 2,6-dichlorophenol 3,4-dichlorophenol 2,4,5-trichlorophenol 2,4,6-trichlorophenol 3,4,5-trichlorophenol 2,3,4,5-tetrachlorophenol 2,3,4,6-tetrachlorophenol pentachlorophenol 4-nitrophenol 2,4-dinitrophenol 2,6-dinitrophenol 3,4-dinitrophenol 2-methyl-4,6-dinitrophenol 4-methyl-2,6-dinitrophenol 2-sec-butyl-4,6-dinitrophenol 2-tert-butyl-4,6-dinitrophenol 4-tert -butyl-2,6-dinitrophenol

24DCP 26DCP 34DCP 245TriCP 246TriCP 345TriCP 2345TeCP 2346TeCP PCP 4NP 24DNP 26DNP 34DNP DNOC DNPC Dinoseb Dino2terb Dino4terb

7.85 6.97 8.59 6.94 6.15 7.73 6.35 5.40 4.75 7.08 3.94 3.70 5.48 4.31 4.06 4.62 4.80 4.11

8.98 8.71 9.61 8.49 7.58 9.27 7.62 6.48 6.13 8.85 4.77 3.81 6.83 4.57 4.15 5.26 5.36 4.69

2.98 2.00 3.30 3.52 2.70 4.13 3.74 2.97 3.32 1.95 1.42 1.32 1.74 1.94 1.74 2.82 3.03 2.69

0.010 0.019 0.002 0.031 0.207 0.005 0.194 0.770 0.880 0.014 0.994 0.999 0.599 0.996 0.999 0.982 0.978 0.995

0.010 0.019 0.002 0.030 0.164 0.005 0.156 0.177 0.106 0.014 0.006 0.001 0.240 0.004 0.001 0.018 0.022 0.005

a All data from ref 13. b Acidity constant in the aqueous phase. c Operational acidity constant in the membrane phase, calculated by eq 13, using data from ref 13. d Distribution ratio between chromatophores and water at pH 7.0 (eq 11). e Fraction of total phenol inside the membrane that is present as phenoxide (eq 14). f Product of the fraction of phenoxide times fraction of neutral species inside the membrane.

TABLE 2

Summary of the Experimental Data Describing the Uncoupling Activity of the Phenols at pH 7.0 compounda

ab

bb

r 2(n)c

24DCP 26DCP 34DCP 245TriCP 246TriCP 345TriCP 2345TeCP 2346TeCP PCP 4NP 24DNP 26DNP 34DNP DNOC DNPC Dinoseb Dino2terb Dino4terb

s.o.g 2.o. s.o. s.o. 12 s.o. 260 43 385 s.o. 260 7 960 350 18 1910 4500 20

510 510 690 58 000 1 540 63 200 1 200 000 15 400 38 900 19 900 f.o.i 3 400 874 000 f.o. 310 f.o. f.o. f.o.

0.996(6) 0.996(4) 0.998(6) 0.963(10) 0.999(8) 0.948(8) 0.951(9) 0.997(11) 0.995(12) 0.996(7) 0.978(15) 0.996(5) 0.997(9) 0.998(6) 0.983(6) 0.992(9) 0.958(5) 0.898(7)

log k1d d.n.p.h d.n.p. d.n.p. d.n.p. 1.8 d.n.p. 3.1 1.7 2.6 d.n.p. 2.4 0.9 3.2 2.5 1.3 3.3 3.7 1.3

log k′2 d

-log C 500ms cph -1)e (mol kgcph

rel importance of 1st-order 500 (%) term in eq 5 at C cph

500ms -log C w (mol L-1)f

4.7 4.4 5.5 6.3 4.0 7.1 6.9 4.9 5.6 6.2 d.n.p. 6.7 6.6 d.n.p. 5.4 d.n.p. d.n.p. d.n.p.

1.3 1.3 1.4 2.3 1.6 2.3 3.0 2.1 2.6 2.1 2.3 1.7 3.1 2.4 1.4 3.1 3.5 1.2

0.95, see Tables 1 and 2). This result is consistent with the assumption that, in these cases, the acceleration of the decay of the membrane potential is caused predominantly by diffusion of A- species through the lipid bilayer. Such “first-order” uncouplers include most of the dinitrophenols with the exception of DNPC and 26DNP, which are very poor first-order uncouplers (see discussion below). For the compounds exhibiting very small f Acph - values (i.e., f Acph - , 0.1), on the other hand, a pure second-order dependence was found that is consistent with the assumption that in these cases AHA- is the important diffusing species. These second-order uncouplers include most of the di- and trichlorophenols as well as 4-nitrophenol (see Tables 1 and 2). Finally, and again consistent with the model, a “mixed-order” dependence of kobs on Ccph was found for those compounds for which at pH 7.0, both the neutral and anionic species are present in appreciable amounts in the chromatophores (i.e., 0.1 < f Acph - < 0.9), thus allowing both A- and AHA- to contribute significantly to the uncoupling effect. 500ms From C cph and the concentration of reaction centers per weight unit of chromatophores (typically approximately 10-7 molRC gcph-1), the number of phenols required for the uncoupling of one reaction center can easily be estimated (7). The number of phenols exceeded the number of reaction centers for all compounds. Consequently, it was not necessary to include the possibility of multiple turnovers of one phenol molecule during one flash experiment into our kinetic model. Effect of pH on kobs. For two of the most potent uncouplers investigated, i.e., Dinoseb and 2345TeCP, some preliminary experiments were conducted to evaluate the

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FIGURE 4. Effect of pH on kobs of (O) 2345TeCP and (2) Dinoseb. The pKacph values of the two compounds are 7.62 and 5.26, respectively (see Table 1).

effect of pH on kobs. Figure 4 shows that, at a fixed total phenol concentration in the chromatophores of 1.5 × 10-3 mol kgcph-1, both compounds showed a maximum in kobs at a solution pH close to their apparent pKacph (see eq 13 and Table 1), that is, at a pH where f Acph - is about 0.5. In the light of our model, changes in kobs caused by pH changes can be explained by two different factors, i.e., (i) changes in the speciation of the phenol inside the membrane and (ii) pH dependence of k1 and k′2. When only considering speciation, it can easily be seen that when eq 4 is rewritten as 2 2 kobs ) -[k′2Ccph ]f Acph2 + [k1Ccph + k′2Ccph ]f Acph (15)

kobs should have a maximum at

f Acph(kobs,max) ) 0.5 +

k1 2k′2Ccph

(16)

For compounds for which k1 , k′2Ccph, a maximum in kobs is expected at or slightly above f Acph - ) 0.5. For 2345TeCP both k1 and k′2 values could be derived at pH 7.0. They are in the order of 103 kgcph mol-1 s-1 and 107 kgcph2 mol-2 s-1 (Table 2). Insertion of these values into eq 15 and with Ccph ) 1.5 × 10-3 mol kgcph-1, a f Acph (kobs,max) of 0.53 is calculated, corresponding to a solution pH of about 7.6, which does not exactly match the experimental data. In fact, a more detailed analysis of the data shows that (also for 2345TeCP) k1 and k′2 cannot be assumed to be constant over the whole pH range investigated and that, therefore, speciation alone cannot solely explain the pH dependence of kobs. The second factor, the pH dependence of the rate constants k1 and k′2 is illustrated by the pH dependence of Dinoseb. Over the pH range of the experiment, k′2 was found to be negligibly small, even at f Acph < 0.5. For reasons discussed below, Dinoseb probably has a very low tendency to form a heterodimer. Hence, kobs would be proportional to f Acph - if k1 was constant over the pH range of the experiment. However, log k1 decreased from 3.5 to 2.7 between pH 5.5 and 9.0 (data not shown). Since k1 describes the mobility of a charged molecule in a membrane with charged head groups, changes in membrane properties due to deprotonation of phospholipid head groups may well cause the observed decrease of k1 with increasing pH.

A more systematic investigation of the effect of pH on the uncoupling activity of phenolic compounds is obviously required and is presently in progress in our laboratory. Additional Structure-Activity Considerations. As has become evident from the above discussion, the intrinsic uncoupling activity of a given phenol (as expressed, for 500ms example, by C cph , see Table 2) depends strongly on its speciation inside the membrane, which is determined by the pKaw of the compound and by the membrane-water partitioning behavior of the phenoxide relative to the neutral species. However, some significant differences in 500ms the C cph values of the compounds investigated are found that cannot be rationalized by speciation only. Obviously, other compound properties, which are reflected in the k1 and/or k′2 values, need to be considered. These properties should include primarily the ability of the phenoxide (or heterodimer) to diffuse through the lipid bilayer as well as the tendency of the phenol to form a heterodimer inside the membrane. The following qualitative analysis of the k1 and k′2 values provides therefore an opportunity to test the model. First, we consider the various dinitrophenols, which, except for 34DNP, are present at pH 7.0 predominantly as phenoxide species. Hence, the large differences (up to 2 orders of magnitude, see Table 2) in their uncoupling activities have to be attributed primarily to differences in the mobility of the respective phenoxides inside the membrane. Qualitatively, it can be expected that the mobility of a phenoxide species in the nonpolar lipid bilayer decreases the more the charge is localized in one part of the molecule, because it can be assumed that with increasing localization of the charge it becomes increasingly difficult to accommodate the molecule in a nonpolar environment. In addition, it is conceivable that phenoxide species with a more localized charge may interact more strongly with the polar groups in the membrane, which would also decrease their mobility. Inspection of the k1 values in Table 2 shows that the data fit this simple picture quite well. All three dinitrophenols exhibiting both nitro groups in ortho-position to the phenoxide group (i.e., 26DNP, DNPC, Dino4terb) are considerably less active uncouplers than the other dinitrophenols where the nitro group in the para-position allows better delocalization of the negative charge over the entire molecule. Charge delocalization may also explain the large difference in the k1 values of 34DNP and 24DNP. Furthermore, among the ortho,para-dinitrophenols, those exhibiting a bulky alkyl substituent in the other ortho-position (i.e., Dinoseb, Dino2terb) are more potent uncouplers than those with no such substituent (i.e., 24DNP, DNOC), while a bulky alkyl substituent in para-position seems to have no significant effect, although it increases significantly the hydrophobicity of the compound (compare Dino4terb with DNOC). These findings can be explained by a shielding of the negative charge at the oxygen by a bulky nonpolar ortho-substituent, which enhances the mobility of the phenoxide (2). The qualitative arguments used to explain the differences in the k1 values of the dinitrophenols can also be extended to the chlorophenols, both when considering the mobility of the phenoxide as well as that of the heterodimer. In addition, for a discussion of the k′2 values, the tendency of a phenol to form a heterodimer has to be taken into account. For the chlorophenols, it is reasonable to postulate that, due to steric interactions, formation of a heterodimer is hindered by chlorine substituents in ortho-position to the

hydroxyl group. Consequently, chlorophenols exhibiting two ortho-Cl should be particularly poor uncouplers for two reasons: localization of the charge as well as steric hindrance with respect to heterodimer formation. Again, the experimental evidence is in accordance with this simple picture. Large differences are found in the k′2 values of isomers, and in all cases, k′2 decreases significantly with the increasing number of ortho-Cl. The same pattern holds for the few k1 values available for the chlorophenols. In this context, it is interesting to compare the k1 and k′2 values of the two tetrachlorophenols with those of pentachlorophenol. For both k1 and k′2, the same sequence (i.e., 2345TeCP > PCP > 2346TeCP) is found. However, the difference between 2346TeCP and the other two compounds is much more pronounced in the case of k′2, which can be rationalized by a steric hindrance of heterodimer formation by the additional ortho-Cl present in 2346TeCP and PCP. A final example that is consistent with the model assumption of heterodimer formation, particularly for compounds that due to their speciation act primarily as second-order uncouplers, is the large difference in uncoupling activity found between 4-nitrophenol (4NP) and 2-nitrophenol (2NP, data not shown). 4NP exhibits a large k′2 value with respect to 2NP (data not shown). Although both compounds exhibit a very similar partitioning behavior, no uncoupling activity could be detected for 2NP even at an aqueous concentration 10-fold higher than the 500ms one corresponding to C cph of 4NP. This result can be explained by the fact that 2NP may form an intramolecular hydrogen bond and therefore, has a much lower tendency to undergo intermolecular hydrogen bonding to form a heterodimer. In conclusion, all the experimental evidence collected in this study supports the shuttle mechanism model described by eq 4. The results also confirm that liposomewater distribution ratios seem to be well suited for estimating the total concentration and speciation of phenolic compounds in biological membranes. To our knowledge, this is the first study in which, for a given class of ionogenic organic compounds, a direct quantitative measure of a specific toxic effect (i.e., uncoupling) could be related to the actual concentration and speciation of the compounds at the target site (i.e., in the membrane).

Ecotoxicological Significance and Outlook From an ecotoxicological point of view, the results of this study are relevant for several reasons. First, with the approach taken, the relative importance of the molecular factors and processes (i.e., uptake, speciation in the membrane, formation of heterodimers, membrane permeability of the charged species) that determine the overall uncoupling potency of a given compound can be assessed. At pH 7.0, this overall uncoupling potency can be expressed 500ms 500ms by C w (see Table 2), which in analogy to C cph is the total phenol concentration in the aqueous phase (in 500ms equilibrium with C cph , see footnote in Table 2), yielding a decay of the membrane potential with a half-life of 500 500ms ms. As shown elsewhere (4), C w values as determined in this study by time-resolved spectroscopy correlate very well with data from various bioassays using other energy500ms transducing systems (21-23). Thus, the C w values should be appropriate measures of the uncoupling potencies of the compounds at pH 7. 500ms In Figure 5, the reciprocal values of C w are plotted 500ms versus the reciprocal values of C cph . Note that in the VOL. 30, NO. 10, 1996 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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500ms 500ms FIGURE 5. Plot of 1/C w versus 1/C cph (log-log scale) for the model compounds (see also Table 2). The solid lines indicate the location of compounds exhibiting Dcphw (pH 7) values of 101, 102, 103, 500ms 500ms and 104 L kg-1cph, respectively (see eq 17). C cph and C w are the equilibrium concentrations in the membrane and in the water, respectively, that cause a decay of the membrane potential with a half-life of 500 ms.

toxicological literature it is common to express toxicity by the reciprocal value of the concentration that causes a 500ms certain effect. Hence, an increasing 1/C w value means 500ms an increasing uncoupling potency. 1/C w is related to 500ms 1/C cph by

1 C

500ms w

1 ) Dcphw(pH 7) 500ms C cph

(17)

From Figure 5, which is a log-log representation of eq 17, the relative contribution of partitioning to the overall uncoupling potency of a given compound at pH 7.0 can be seen immediately. It is, for example, interesting to note that the large difference in uncoupling potency between 345TriCP and 24DNP (factor 500) is entirely due to differences in the partitioning of the two compounds, while the similarly large difference between Dino2terb and Dino4terb has to be attributed primarily to the very different mobility of the respective phenoxide species in the membrane (see discussion above). Furthermore, when com500ms paring compounds with similar 1/C w values, it can be seen that in most cases the contribution of partitioning is much larger in the case of the chlorophenols as compared to the less hydrophobic nitrophenols, which exhibit larger 500ms 1/C cph values due to their lower pKacph and more even charge distribution within the molecule. Finally, as is also immediately evident from Figure 5, adding a bulky alkyl substituent, such as a tertiary butyl group, in ortho-position to the phenolic group may increase the uncoupling potency by several orders of magnitude (compare Dino2terb with 24DNP) by enhancing both the partitioning into as well as the mobility of the phenoxide in the membrane. In the case of Dino2terb, the two contributions are of about similar magnitude. Hence, one could, for example, speculate that substitution of 34DNP with a tert-butyl group in orthoposition to the OH group would create a compound that should exhibit a significantly higher uncoupling potency than Dino2terb. These examples demonstrate that the approach taken in this study allows one to gain important insights into the

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factors that determine the uncoupling potency of phenolic compounds in aquatic systems. The results should contribute significantly to a better understanding of observed toxic effects of phenolic compounds. They also form a base for the development of more powerful quantitative structure-activity relationships than those presently available (e.g., refs 21 and 23). To this end, however, the set of model compounds needs to be extended, and the effect of pH on the uncoupling potency of phenolic compounds has to be investigated more carefully. This is the topic of an ongoing study in our laboratory. In addition to contributing to a better assessment of the uncoupling potencies of phenolic compounds, this study also offers other interesting perspectives with respect to future work. One very important topic that needs to be addressed more extensively in the future is the toxic effect of mixtures of organic compounds on organisms. The method presented in this paper should be very well-suited to study synergistic and antagonistic effects on energy transduction of phenolic compounds when present in mixtures. For example, it can be checked as to what extent uncoupling acitivities are additive, or whether antagonistic effects (e.g., by competition for sites in the membrane) may play a significant role. Furthermore, the possible enhancement of the uncoupling activity of a first-order uncoupler by additional heterodimer formation with a hydrophobic neutral phenol also present in the membrane can be studied. Such work is presently also in progress. Finally, the approach taken in this study should also be applicable to other classes of compounds that exhibit uncoupling properties, including, for example, diarylamines (24), aromatic amines (25) and organotin compounds (26).

Acknowledgments We are indebted to John Westall for many helpful discussions and to Tim Grundl, Stefan Haderlein, Jo¨rg Klausen, and John Westall for reviewing the manuscript. Valuable comments were made by Renata Behra and Hauke Harms. This project was supported by a grant of the Swiss Federal Institute of Technology (TH Project 27.92-1).

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(14) Benz, R.; McLaughlin, S. Biophys. J. 1983, 41, 381-398. (15) Kasianowicz, J.; Benz, R.; McLaughlin, S. J. Membr. Biol. 1984, 82, 179-190. (16) Finkelstein, A. Biochim. Biophys. Acta 1970, 205, 1-6. (17) Ba¨uerle, H.; Seelig, J. Biochemistry 1991, 30, 7203-7211. (18) Dutton, P. L.; Petty, K. M.; Bonner, H. S.; Morse, S. D. Biochim. Biophys. Acta 1975, 387, 536-556. (19) Petitou, M.; Tuy, F.; Rosenfeld, C. Anal. Biochem. 1978, 91, 350353. (20) Junge, W.; Jackson, J. B. In Photosynthesis: Energy Conversion by Plants and Bacteria; Govindjee, Ed.; Academic Press: New York, 1982; Vol. 1, pp 589-646. (21) Miyoshi, H.; Tsujishita, H.; Tokutake, N.; Fujita, T. Biochim. Biophys. Acta 1990, 1016, 99-106. (22) Argese, E.; Bettiol, C.; Ghelli, A.; Todeschini, R.; Miani, P. Environ. Toxicol. Chem. 1995, 14, 363-368.

(23) Ravanel, P.; Taillandier, G.; Tissut, M. Ecotoxicol. Environ. Saf. 1989, 18, 337-345. (24) Guo, Z.; Miyoshi, H.; Komyoji, T.; Haga, T.; Fujita, T. Biochim. Biophys. Acta 1991, 1059, 91-98. (25) Schultz, T. W.; Lin, D. T.; Arnold, M. Sci. Total Environ. 1991, 109/110, 569-580. (26) Fent, K. Crit. Rev. Toxicol. 1996, 26, 1-117. (27) Melandri, B. A.; Mehlhorn, R. J.; Packer, L. Arch. Biochem. Biophys. 1984, 235, 97-105.

Received for review February 19, 1996. Revised manuscript received May 14, 1996. Accepted May 15, 1996.X ES960153F X

Abstract published in Advance ACS Abstracts, July 15, 1996.

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