J. Phys. Chem. B 2010, 114, 457–462
457
Solvent Properties Governing Solute Partitioning in Polymer/Polymer Aqueous Two-Phase Systems: Nonionic Compounds Pedro P. Madeira,*,†,‡,§ Celso A. Reis,‡,| Alı´rio E. Rodrigues,† Larissa M. Mikheeva,§ and Boris Y. Zaslavsky§ Laboratory of Separation and Reaction Engineering, Departamento de Engenharia Quı´mica, Faculdade de Engenharia da UniVersidade do Porto, and Institute of Molecular Pathology and Immunology of the UniVersity of Porto, Rua Dr. Roberto Frias s/n, 4200-465 Porto, Portugal, Medical Faculty of the UniVersity of Porto, Alameda Prof. Hernaˆni Monteiro, 4200-319, Porto, Portugal, and Analiza, Inc. 3615 Superior AVenue, CleVeland, Ohio 44114 ReceiVed: July 31, 2009; ReVised Manuscript ReceiVed: October 8, 2009
The solvatochromic solvent parameters characterizing the solvent polarity (π*), solvent hydrogen-bond donor acidity (R), and solvent hydrogen-bond acceptor basicity (β) of aqueous media were measured in the coexisting phases of nine different aqueous polymer/polymer two-phase systems (ATPS), containing 0.15 M NaCl in 0.01 M phosphate buffer, pH 7.4. Partitioning coefficients of six neutral compounds were measured in the nine ATPS at particular polymer concentrations. The solvatochromic equation was used to describe the partitioning of each compound. Three descriptors of the solvent properties of the phases could describe adequately the partitioning of the solutes in all the ATPS employed. 1. Introduction Aqueous two-phase systems (ATPS) arise in aqueous mixtures of different water-soluble polymers or a single polymer and a specific salt. When two specific polymers, for example, dextran and poly(ethylene glycol) (PEG), are mixed in water above certain concentrations, the mixture separates into two immiscible aqueous phases. There is a clear interfacial boundary separating two distinct aqueous-based phases, each preferentially rich in one of the polymers, with the aqueous solvent in both phases providing media suitable for biological products.1-4 These systems are unique in that each of the phases contains over 80% water on a molal basis and yet they are immiscible and differ in their solvent properties;1-5 therefore these systems can be used for differential distribution of solutes and particles. Extraction in ATPS has been clearly demonstrated as an efficient method for large-scale recovery and purification of biological products.1-3,6,7 Low cost, high capacity, and easy scale-up are clear advantages of this technology. Partitioning in ATPS may also be used for characterization of protein surface properties,4,8 changes in protein structure,9 conformation,10 and ligand binding.1-3 For successful utilization of partitioning in ATPS, it is important to understand the mechanisms of solute distribution in the systems as well as system properties at the molecular level. The underlying concept for one current explanation for partitioning in ATPS is that the polymers engaged in the formation of an ATPS are essentially neutral to the solute being partitioned and are important only in regard to their effects on the solvent features of the aqueous media in the coexisting * Corresponding author: tel (+351) 225081578; fax (+351) 225081674; e-mail
[email protected]. Permanent address: Laboratory of Separation and Reaction Engineering, Universidade do Porto. † Laboratory of Separation and Reaction Engineering, Dpt. de Engenharia Quı´mica, Faculdade de Engenharia da Universidade do Porto. ‡ Institute of Molecular Pathology and Immunology of the University of Porto. § Analiza, Inc. | Medical Faculty of the University of Porto.
phases. This view is supported by experimental evidence, which indicates that (a) the solvent features of the aqueous media in the coexisting phases are different4,5 and (b) there are clear similarities between partitioning of solutes in ATPS and in water-organic solvent systems.4,5,11-15 It should be noted that this concept is not applicable to ATPS containing charged polymers or polymers with ligands for so-called affinity partitioning. The Kamlet-Taft solvatochromic comparison approach16-18 was used to study dipolarity/polarizability (π*), hydrogen-bond acceptor basicity (β), and hydrogen-bond donor acidity (R) of aqueous media in the coexisting phases of different two-polymer ATPS. This method has been widely used to measure the properties of solvents and solvent mixtures19-23 including aqueous PEG solutions24,25 and phases of PEG-salt biphasic systems.26 We examined partitioning of a series of nonionic organic compounds in the ATPS characterized with solvatochromic probes and described partitioning of nonionic compounds in the ATPS in terms of solute-solvent interactions. Selection of ATPS was based on the use of all possible combinations of pairs of nonionic polymers on hand, and watersoluble, chromophore-containing organic compounds noncharged at pH 7.4 were selected for partitioning. 2. Experimental Section 2.1. Materials. 2.1.1. Polymers. All polymers were used without further purification. Dextran 75 (lot 115195), weightaverage molecular weight (Mw) = 60000-90000, was purchased from USB (Cleveland, OH). Poly(ethylene glycol) 8000 (lot 069H00341), Mw ) 8000, was purchased from Sigma-Aldrich (St. Louis, MO). A random copolymer of ethylene glycol and propylene glycol, Ucon 50-HB-5100 (lot SJ1955S3D2), Mw ) 3930, was purchased from Dow Chemical (Midland, MI). Ficoll 70 (lot 302970), Mw = 70 000 was purchased from GE Healthcare Biosciences AB (Sweden). Hydroxypropyl starch, Reppal PES-100 (lot D702-09/01), Mw = 100 000, was purchased from Reppe AB (Sweden).
10.1021/jp907346s 2010 American Chemical Society Published on Web 12/18/2009
458
J. Phys. Chem. B, Vol. 114, No. 1, 2010
Madeira et al.
TABLE 1: Polymer Compositionsa of Phases in the Aqueous Two-Phase Systems Used for Partitioningb total composition
top phase
bottom phase
polymer 1
polymer 2
polymer 1
polymer 2
polymer 1
polymer 2
polymer 1
polymer 2
Eb
dextran dextran dextran PES PEG Ficoll Ficoll PES PES
Ficoll PEG Ucon dextran Ucon PEG Ucon Ficoll PEG
12.94 12.41 12.39 17.30 15.00 15.06 13.01 17.31 15.24
18.06 6.06 10.08 12.43 29.97 7.90 9.93 14.86 6.96
3.23 0.31 0.16 5.31 0.36 9.55 2.90 10.31 3.67
28.31 13.02 18.30 21.68 50.25 11.65 16.42 20.20 12.28
21.57 22.44 26.51 31.38 35.46 23.97 24.50 25.35 29.58
9.03 0.53 0.59 1.93 1.58 1.83 2.54 7.80 0.37
0.0191 ( 0.0006 0.0271 ( 0.0008 0.085 ( 0.002 -0.02205 ( (4 × 10-5) 0.123 ( 0.008 0.0097 ( 0.0009 0.05 ( 0.02 -0.0126 ( 0.0007 -0.0147 ( (6 × 10-5)
a Polymer concentrations are given in weight percent. b Parameter E denotes the difference between the relative hydrophobic character of the coexisting phases. It was calculated from experimental data on partitioning of sodium salts of dinitrophenylamino acids with aliphatic side chains described by eq 6 as reported in ref 27; see text for physical meaning of parameter E.
2.1.2. SolWatochromic Dyes. The solvatochromic probes 4-nitrophenol (reagent grade, >98%) and 4-nitroanisole (>97%, GC) were both supplied by Aldrich, Milwaukee, WI. Reichardt’s carboxylated betaine dye, 2,6-dipenyl-4-[2,6-diphenyl-4-(4carboxyphenyl)-1-pyridino]phenolate, sodium salt was kindly provided by Professor C. Reichardt (Philipps University, Marburg, Germany). 2.1.3. Other Chemicals. All salts and other chemicals used were of analytical-reagent grade. 2.2. Methods. 2.2.1. SolWatochromic Studies. ATPS of the compositions listed in Table 1 were prepared as previously described.27,28 The phases were separated and used for solvatochromic analysis. The solvatochromic probes 4-nitroanisole (ca. 10 mM), 4-nitrophenol (ca. 10 mM), and Reichardt’s carboxylated betaine (ca. 10 mM) were used to measure the polarity/polarizability π*, H-bond acceptor (HBA) basicities β, and H-bond donor (HBD) acidities R in both phases of each particular ATPS. Three to four concentrations of each probe were prepared for each analysis in order to check for aggregation effects and specific interactions with the phase-forming polymers by adding 10-40 µL of solvatochromic probe stock solution to 1.1 mL of the ATPS phase sample. Aliquots (20 µL) of 1 M NaOH were added to the samples containing Reichardt’s carboxylated betaine to ensure pH > 8.0. The samples were mixed thoroughly in a vortex mixer and then scanned in a UV-vis spectrophotometer (Thermo Electron Corp., model UV1) with a bandwidth of 2.0 nm; data interval 0.2 nm; and Hi-res scan (∼0.5 nm/segment). Pure phases containing no dye (blank) were scanned first to establish a baseline. Identical phases containing dye additives were subsequently scanned in the UV-vis spectrophotometer in triplicate or more. The maximum wavelength was the average between these scans. Average standard deviation for each measured wavelength was below 1.0 nm and in most cases e0.7 nm for all probes. Water (20 µL) was added to the systems to prevent clouding. A few drops of HCl (1 M) were added to the samples containing 4-nitrophenol in order to eliminate charge transfer bands that were observed in some systems. It should be noted that the wavelength of maximum absorbance was determined for 4-nitrophenol and for Reichardt’s carboxylated betaine dye in several solvents (e.g., water, n-hexane, methanol) and phases of ATPS in the presence and absence of HCl (for 4-nitrophenol) and/or NaOH (for the betaine dye) at different concentrations of the probes as well as different concentrations of HCl or NaOH. The solvatochromic shift of a probe in a given medium in all cases was within the experimental error (data not shown).
a. Determination of SolVent Dipolarity/Polarizability π*. π* was determined from the wavenumber (ν1) of the longest wavelength absorption band of the 4-nitroanisole dye by use of the relationship given by Marcus:29
π* ) 0.427(34.12 - ν1)
(1)
b. Determination of SolVent Hydrogen-Bond Acceptor Basicity β. β values were determined from the wavenumber (ν2) of the longest wavelength absorption band of the 4-nitrophenol dye by use of the relationship29
β ) 0.346(35.045 - ν2) - 0.547π*
(2)
c. Determination of SolVent Hydrogen-Bond Donor Acidity R. R values were determined from the longest wavelength absorption band of Reichardt’s betaine dye by use of the relationship29
R ) 0.0649ET(30) - 2.03 - 0.72π*
(3)
The empirical Reichardt’s solvent polarity index, ET(30) value, is obtained directly from the wavelength absorption band of the probe as
ET(30) ) 1/0.932(28 591/λ - 3.335)
(4)
where λ is the wavelength in nanometers. 2.2.2. Partitioning. a. Phase Systems. A mixture of polymers was prepared by dispensing appropriate amounts of the aqueous stock polymer solutions into a 1.2 mL microtube by use of a Hamilton Company (Reno, NV) ML-4000 four-probe liquid handling workstation. Appropriate amounts of stock buffer solutions were added to give the required ionic and polymer composition of the final system with a total weight of 0.5 g (after addition of sample solution, see below). All two-phase systems had the polymer compositions indicated in Table 1 and salt composition of 0.15 M NaCl in 0.01 M sodium phosphate buffer (NaPB), pH 7.4. b. Partition Experiments. Partition coefficient for each nonionic organic compound was measured in a set of ATPS of the same composition with five different amounts of a given compound varied over about 1-1.5 orders of magnitude range. An automated instrument for performing aqueous two-phase partitioning, Automated Signature Workstation, ASW (Analiza, Cleveland, OH), was used for the partition experiments. The ASW system is based on the ML-4000 liquid-handling workstation (Hamilton Company, Reno, NV) integrated with a FL600 fluorescence microplate reader (Bio-Tek Instruments, Winooski, VT) and a UV-vis microplate spectrophotometer (SpectraMax
Solvent Properties Governing Solute Partitioning
J. Phys. Chem. B, Vol. 114, No. 1, 2010 459
TABLE 2: Solvatochromic Solvent Parametersa and Their Differencesb in the Coexisting Phases of Aqueous Two-Phase Systems top phase
bottom phase
no.
polymer 1c
polymer 2c
π*
R
β
π*
R
β
∆π*b
∆Rb
∆βb
1 2 3 4 5 6 7 8 9
dextran dextran dextran PES PEG Ficoll Ficoll PES PES
Ficoll PEG Ucon dextran Ucon PEG Ucon Ficoll PEG
1.188 1.099 1.112 1.275 1.035 1.116 1.146 1.137 1.116
0.984 1.078 0.882 0.995 0.628 0.999 0.850 1.050 1.032
0.633 0.632 0.659 0.596 0.757 0.612 0.667 0.668 0.595
1.150 1.167 1.179 1.163 1.158 1.167 1.031 1.196 1.175
1.048 1.096 1.023 0.997 0.766 0.976 1.063 0.996 0.962
0.678 0.624 0.597 0.681 0.697 0.634 0.644 0.628 0.697
0.038 -0.068 -0.068 0.113 -0.123 -0.051 0.115 -0.059 -0.059
-0.064 -0.018 -0.141 -0.002 -0.138 0.023 -0.213 0.054 0.069
-0.045 0.008 0.062 -0.084 0.060 -0.022 0.023 0.040 -0.102
a π*, solvent dipolarity/polarizability; R, solvent hydrogen-bond donor acidity; β, solvent hydrogen-bond acceptor basicity. b All differences are calculated between values measured in the top phases and those measured in the bottom phases. c Polymer 1, predominant polymer in the bottom phase; polymer 2, predominant polymer in the top phase (for polymer composition of each phase, see Table 1).
Plus384, Molecular Devices, Sunnyvale, CA). Solutions of all compounds to be partitioned were prepared in water at concentrations of 1-5 mg/mL. Varied amounts (e.g., 0, 15, 30, 45, 60, and 75 µL) of a given compound solution and the corresponding amounts (e.g., 100, 85, 70, 55, 40, and 25 µL) of water were added to a set of the same polymer/buffer mixtures. Systems were vortexed in a Multipulse vortexer and centrifuged for 30-60 min at 3000g in a refrigerated centrifuge (Jouan, BR4i, Thermo Fisher Scientific, Waltham, MA) to accelerate phase settling. The upper phase in each system was removed, the interface was discarded, and aliquots of 20-70 µL from the upper and lower phases were withdrawn in duplicate for analysis. Aliquots from both phases were diluted with water up to 320 µL in microplate wells. Following moderate shaking for 20 min in an incubator (Vortemp 56EVC, Labnet International, Edison, NJ) at room temperature (23 °C), the UV-vis plate reader was used to measure optical absorbance at wavelengths previously determined to correspond to maximum absorption. Phases of blank systems at corresponding dilutions were measured for comparison. The partition coefficient K is defined as the ratio of the sample concentration in the upper phase to the sample concentration in the lower phase. The K value for each solute was determined as the slope of the concentration in the upper phase plotted as a function of the concentration in the bottom phase averaged over the results obtained from 2-4 partition experiments carried out at the fixed composition of the system. The advantages of this procedure over more common protocol, including several partition experiments of a solute at the same concentration, are discussed in detail elsewhere (ref 4, pp 223-226). Deviation from the average K value was consistently below 5% and in most cases lower than 2%. 3. Results and Discussion It should be noted that each of the solvent parameters π*, β, and R were obtained from a set of single solvatochromic probes as described above instead of using the average of results from several multiple probes as suggested by Kamlet et al.19 The use of a single probe versus multiple probes remains an issue debated in the literature.26,29,30 Huddleston et al.26 demonstrated that using a set of single solvatochromic probes provide reasonable results for analysis of solvent features of coexisting phases of PEG-salt ATPS. The solvatochromic parameters measured in each phase of the ATPS (see Table 1) are presented in Table 2. The differences between the values found for the top phases and those for the corresponding bottom phases are also shown. It should be
mentioned that dipolarity/polarizability, π*, of pure water is 1.091 and that of the salt solution (0.15 M NaCl in 0.01 M sodium phosphate buffer, pH 7.4) is 1.097. It can be seen from the data presented in Table 2 that the dipolarity/polarizability is unequal between coexisting phases for each system and most values are slightly higher than that of pure water and the background salt solution. The exception to this was the Uconrich top phase in the PEG-Ucon ATPS. From observation with a N,N-diethyl-4-nitroaniline probe, Singh and Pandey25 reported a decrease in π* with PEG concentration in water, though Kim et al.24 tested five different solvatochromic probes and observed no change in dipolarity/polarizability relative to water. Huddleston et al.26 reported a very slight increase in π* value in the PEG-salt ATPS phases as measured with 4-nitroanisole dye. It is possible that the difference between the dipolarity/ polarizability of the coexisting phases is a function of the concentration of the second polymer (dextran, PES, Ficoll, or Ucon) rather than PEG. The largest difference in dipolarity/ polarizability between the coexisting phases was observed in the PEG-Ucon, Ficoll-Ucon, and PES-dextran systems. The solvent hydrogen-bond acidity, R, in the buffer solution was found to be 1.271, while it was reported as 1.17 in pure water. The solvent acidity measured in each phase was consistently lower than that of buffer solution, though R values differed between coexisting phases of each system. Ficoll-Ucon and PEG-Ucon are among those with the largest difference. The dextran-Ucon system has significant ∆R while pretty average ∆π*, and the coexisting phases of the aforementioned PES-dextran system are essentially similar in regard to solvent hydrogen-bond donor acidity. Data reported for aqueous PEG solutions24,25 and phases of PEG-salt ATPS26 suggests that hydrogen-bond acidity decreases with increasing PEG concentration. In contrast, hydrogen-bond basicity, β, reportedly increased with PEG concentration and was also higher in the PEG-rich phase relative to the salt-rich phase in PEG-salt ATPS. Solvent basicity, β, of the buffer solution was determined to be 0.611, that is, slightly higher than that reported for pure water (0.606). It can be seen from the data in Table 2 that the solvent basicity is increased or slightly reduced depending on the phaseforming polymers. In all the ATPS under study, the solvent hydrogen-bond acidity and basicity values in the coexisting phases showed no correlation, which can be readily seen from the signs of the corresponding ∆R and ∆β values in Table 2. According to the Kamlet-Taft solvatochromic comparison approach,16-18 the nonionic solute partition coefficient may be expressed as31
460
J. Phys. Chem. B, Vol. 114, No. 1, 2010
Madeira et al.
log Ks ) z0 + ss∆π* + bs∆β + as∆R + Vs∆δH
(5) where K is the solute partition coefficient; π*, R, and β are as defined above; the δH term is the Hildebrand solubility parameter, which serves as a measure of the solvent/solvent interactions that are interrupted in creating a cavity for the solute;19 s, a, and b are coefficients that measure the relative susceptibilities of the partition coefficient to the indicated solvent properties; Vs is the solute molecular volume; the subscript s designates the solute; and ∆ denotes the difference for a given solvent property between the coexisting phases. Each ATPS under study was characterized earlier27 in terms of the free energy of transfer of a CH2 group between the coexisting phases. This approach4,5,11,27,32-37 is based on the partitioning of a homologous series of solutes with varied aliphatic alkyl chain length in a given two-phase system described as
log Kj ) c + E[N(CH2)j]
(7)
where ln K is the natural logarithm of the partition coefficient K, R is the universal gas constant, and T is the absolute temperature in kelvins. It follows that
∆G(CH2) ) -RTE
δH2 ) 0.229((0.0096) + 0.212((0.018)π* N ) 23
R ) 0.8493 2
SD ) 0.0297
(8)
where ∆G(CH2) is the free energy of transfer of a methylene group between the coexisting phases. The free energy of transfer of a methylene group between the phases is generally considered4,5,11 to be approximately equivalent to the relative free energy difference in cavity formation between the phases. In aqueous organic solvent systems the relative free energy of cavity formation between two phases is represented19,31 by the δH term (Hildebrand solubility parameter) as shown above in eq 5. Analysis of the data presented by Marcus31 for biphasic systems formed by water-immiscible essentially dry organic solvents and water shows that for 23 solvents there is a relationship between the solvent cohesive energy density (square of the Hildebrand
(9)
F ) 118.4
N is the number of solvents; F is the ratio of variance, SD is the standard deviation, and R2 is the correlation coefficient (data for water, bromoform, and di-2-propyl ether were excluded from analysis). Discussion of the relationship described by eq 9 is beyond the scope of the present work. Analysis of the relative free energy difference in cavity formation between the phases of ATPS (parameter E in eq 6) in terms of solvatochromic parameters showed that there is a relationship between the coefficients E presented in Table 1 and parameters ∆π* and ∆R, presented in Table 2, described as E ) -0.001((0.004) - 0.34((0.05)∆π* - 0.46((0.04)∆R
(6)
where log Kj is the logarithm of the distribution coefficient K of a jth member of the homologous series with the corresponding N(CH2)j length of the aliphatic chain of a given solute. E is an average log K increment per CH2 group; and c represents the total contribution of the non-alkyl part of the structure of the solute in the series into log Kj. The free energy of transfer of a solute between the phases is given by eq 7:
∆G ) -RT ln K
solubility parameter, δH2) and the solvent solvatochromic polarity/polarizability:
(10) N)9
R ) 0.9620 2
SD ) 0.011
F ) 75.9
N is the number of systems, F is the ratio of variance, SD is the standard deviation, and R2 is the correlation coefficient. The relationship described by eq 10 indicates that the relative free energy difference in cavity formation between the phases of ATPS depends on both solvent polarity/polarizability, π*, and solvent hydrogen-bond acidity, R, of the aqueous media in the coexisting phases. It seems reasonable that the free energy of cavity formation in an aqueous medium, resulting in rearrangement of the highly cooperative hydrogen-bonds network, would involve all types of solvent-solvent interactions. It is unclear why the hydrogen-bond basicity of aqueous media is not a factor in the free energy of cavity formation. Partition coefficients of nonionic compounds examined in each ATPS are presented in Table 3. Analysis of the data in terms of parameters ∆π*, ∆R, and ∆β, presented in Table 2, shows an existence of the linear correlations for all these compounds expressed as:
log Ks ) z0 + ss∆π* + as∆R + bs∆β
(11)
where all the parameters are as defined above. The regression coefficients ss, as, and bs determined by multiple linear regression of the solvent parameters on a logarithm of the solute partition coefficient in each ATPS are presented in Table 4. The partition coefficients for each of the nonionic compounds in different ATPS coincided sufficiently with the solvatochromic solvent parameters measured. The
TABLE 3: Partition Coefficients K Determined for Nonionic Solutes in the ATPS Indicateda solute
DexFicoll
DexPEG
PEGUcon
DexUcon
PESPEG
FicollPEG
FicollUcon
PESDex
PESFicoll
benzyl alcohol 3-hydroxybenzaldehyde nitrobenzene phenol 4-hydroxyacetanilide caffeine 4-NP-β-D-fucopyranoside 4-NP-β-D-galactopyranoside 4-NP-β-D-glucopyranoside 4-NP-R-D-glucopyranoside 4-NP-R-D-mannopyranoside
1.18 1.31 0.94 1.28 1.32 1.09 1.22 1.25 1.22 1.18 1.23
1.64 1.74 0.98 1.90 2.12 1.15 1.31 1.18 1.27 1.23 1.18
2.58 4.74 5.81b 4.00 2.56b 1.55 2.74 2.54 2.54 2.55 3.14
2.05 3.29 1.77 3.04 3.68 1.43 1.55b 1.40b 1.54b 1.59b 1.60b
1.04 1.10 0.56 1.17 1.26 0.93 0.87 0.81 0.86 0.86 0.77
1.17 1.16 0.70 1.21 1.30 1.01 1.02 0.99 1.00 1.02 0.98
1.52 2.13 1.22 2.02 2.21 1.24 1.24 1.16 1.25 1.27 1.28
0.72 1.52b 0.45 0.72 0.66 0.84 0.76 0.74 0.78 0.78 0.70
1.94 0.94 0.60 0.95 0.90 0.96 0.91 0.90 0.92 0.92 0.85
a
K values for 4-nitrophenyl glycopyranosides are presented here as reported previously.27,28 correlation described by eq 11.
b
Partition coefficient values not fitting the
Solvent Properties Governing Solute Partitioning
J. Phys. Chem. B, Vol. 114, No. 1, 2010 461
TABLE 4: Regression Coefficients in Equation 11a solute
z0
benzyl alcohol 0.14 ( 0.03 3-hydroxybenzaldehyde 0.00 ( 0.01 nitrobenzene -0.18( 0.02 phenol 0.03 ( 0.01 4-hydroxyacetanilide 0.02 ( 0.02 caffeine -0.007 ( 0.003 p-NP-β-D-fucopyranoside -0.02 ( 0.03 p-NP-β-D-galactopyranoside -0.03 ( 0.04 p-NP-β-D-glucopyranoside -0.02 ( 0.03 p-NP-R-D-glucopyranoside -0.02 ( 0.03 p-NP-R-D-mannopyranoside -0.05 ( 0.04 a
ss
as
bs
-0.8 ( 0.3 -0.3 ( 0.3 1.9 ( 0.5 -2.5 ( 0.1 -2.9 ( 0.1 -0.8 ( 0.2 -1.7 ( 0.3 -2.3 ( 0.3 -0.2 ( 0.4 -2.2 ( 0.2 -2.5 ( 0.2 -0.8 ( 0.3 -2.9 ( 0.3 -3.1 ( 0.3 -1.2 ( 0.4 -0.65 ( 0.04 -0.87 ( 0.04 -0.06 ( 0.07 -1.6 ( 0.3 -1.7 ( 0.3 -0.4 ( 0.6 -1.5 ( 0.4 -1.6 ( 0.4 -0.2 ( 0.7 -1.5 ( 0.3 -1.6 ( 0.3 -0.3 ( 0.5 -1.4 ( 0.3 -1.6 ( 0.3 -0.2( 0.5 -1.8 ( 0.4 -2.1 ( 0.4 -0.3 ( 0.7
N
F
SD
r2
9 8 8 9 8 9 8 8 8 8 8
31 369 59 142 70 381 20 13 19 22 17
0.05 0.02 0.04 0.03 0.04 0.007 0.06 0.07 0.05 0.05 0.07
0.949 0.996 0.978 0.988 0.981 0.996 0.938 0.904 0.934 0.943 0.926
outlierb PES-dex PEG-Ucon PEG-Ucon dex-Ucon dex-Ucon dex-Ucon Dex-Ucon Dex-Ucon
Log Ks ) z0 + ss∆π* + as∆R + bs∆β for indicated compounds. b Outlier: an ATPS in which the K value for a given solute does not fit eq
11.
Figure 1. Partition coefficients K, experimentally measured for all the examined compounds (see Table 3) in each ATPS (Table 1), plotted against K values calculated with eq 11 by use of measured solvatochromic solvent parameters (Table 2) and calculated regression coefficients (presented in Table 4).
regression coefficients determined for each compound represent the relative susceptibilities of the compound partition coefficient to the corresponding solvent properties. The regression coefficients determined for several glycosides28 are also presented in Table 4. K values calculated with eq 11, using regression coefficients listed in Table 4 and solvent parameters in Table 2, agree well with the experimentally determined partition coefficients for all the solutes in each ATPS (Table 3) as shown in Figure 1. It should be noted that the outliers denoted in Table 3 were excluded from calculations of the regression coefficients. Analysis of the regression coefficients in Table 4 indicates that different solute-solvent interactions affect solute partitioning in ATPS differently depending on the particular solute structure. Analysis of the relative contributions of these interactions demonstrates that the solvent hydrogen-bond acidity and basicity of the phases is as important as the dipolarity/ polarizability of the coexisting phases, though it seems to depend upon the particular solute under study. For benzyl alcohol, for example, the difference between the hydrogen-bond solvent basicity of the coexisting phases is more important than solvent acidity, in contrast to caffeine, nitrobenzene, and all the glycosides examined thus far. For partitioning of other compounds it appears that both solvent acidity and solvent basicity are important. Our data in this regard contradict the conclusion drawn by Moody et al.11 that, in an aqueous dextran-PEG twophase system, the hydrogen-bond solvent basicity of the phases plays no role in the partitioning of nonionic compounds. The
results presented in Table 2 indicate the difference between the solvent basicity of the phases in the dextran-PEG ATPS is extremely small, and this may explain the seeming contradiction. It should also be mentioned that according to Moody et al.11 the molecular volume of the partitioned solute affects the solute partition behavior almost as much as the solvent hydrogen-bond acidity. The correlations established in this study do not include the effect of the solute volume. This study examined the relationship between partition coefficients for a given solute in different ATPS and the solvatochromic parameters characterizing the solvent properties of the coexisting phases of these ATPS. The values for the regression coefficients that characterize these relationships for different solutes (Table 4) are likely to include contributions of the factors related to the solute molecular volume effect on the solute partition coefficient. The relationship described by eq 10 supports this assumption. The fact that the relative free energy of cavity formation in the phases of ATPS is related to the solvent polarity/polarizability and solvent hydrogen-bond acidity implies that the effect of the solute molecular volume on the solute partition coefficient may impact the values of the regression coefficients ss, as, and bs in Table 4. The number of solutes examined in this study and the range of their molecular volumes is too limited, however, for exploring this assumption further. The solvatochromic comparison approach used in this work may potentially be used for analysis of partition behavior of biological macromolecules, such as proteins, for which ATPS provide media suitable for separation and analysis. Since these solutes are generally charged in aqueous media, it is expected that an additional term taking into account ion-ion and ion-dipole solute-solvent interactions must be included in the linear solvation energy relationship (LSER) equation. This work is currently in progress. 4. Conclusions The Kamlet-Taft solvatochromic comparison approach was used to characterize dipolarity/polarizability (π*), hydrogenbond acceptor basicity (β), and hydrogen-bond donor acidity (R) of aqueous media in the coexisting phases of nine different two-polymer two-phase systems. The partition coefficients of six nonionic compounds were measured in each system. It was found that the solute partition coefficient can be described by a linear combination of the established solvatochromic parameters of solvent properties in each of the coexisting phases. Acknowledgment. We thank Professor M. H. Abraham (University College London) for helpful discussions and for reviewing the manuscript. PPM acknowledges Prof. Eugénia
462
J. Phys. Chem. B, Vol. 114, No. 1, 2010
Macedo (LSRE-Porto) for the use of her equipment to measure solvatochromic data. P.P.M. acknowledges financial support (Grant SFRH/BPD/45055/2008) from Fundac¸a˜o para a Cieˆncia e a Tecnologia (FCT), Lisbon, Portugal. References and Notes (1) Albertsson, P. A. Partition of Cell Particles and Macromolecules, 3rd ed.; Wiley: New York, 1986. (2) Walter, H.; Brooks, D. E.; Fisher, D. Partitioning in Aqueous TwoPhase Systems: Theory, Methods, Use, and Applications to Biotechnology; Academic Press: Orlando, FL, 1985. (3) Walter, H.; Johansson, G. Methods Enzymol. 1994, 228, XV-XVI. (4) Zaslavsky, B. Y. Aqueous Two-Phase Partitioning: Physical Chemistry and Bioanalytical Applications; Marcel Dekker: New York, 1994. (5) Willauer, H. D.; Huddleston, J. G.; Rogers, R. D. Ind. Eng. Chem. Res. 2002, 41 (11), 2591–2601. (6) Frerix, A.; Geilenkirchen, P.; Mu¨ller, M.; Kula, M. R.; Hubbuch, J. Biotechnol. Bioeng. 2007, 96 (1), 57–66. (7) Persson, J.; Andersen, D. C.; Lester, P. M. Biotechnol. Bioeng. 2005, 90 (4), 442–451. (8) Berggren, K.; Wolf, A.; Asenjo, J. A.; Andrews, B. A.; Tjerneld, F. Biochim. Biophys. Acta 2002, 1596 (2), 253–268. (9) Zaslavsky, A.; Gulyaeva, N.; Chait, A.; Zaslavsky, B. Anal. Biochem. 2001, 296 (2), 262–269. (10) Jensen, P. E. H.; Stigbrand, T.; Shanbhag, V. P. J. Chromatogr., A 1994, 668 (1), 101–106. (11) Moody, M. L.; Willauer, H. D.; Griffin, S. T.; Huddleston, J. G.; Rogers, R. D. Ind. Eng. Chem. Res. 2005, 44 (10), 3749–3760. (12) Willauer, H. D.; Huddleston, J. G.; Rogers, R. D. Ind. Eng. Chem. Res. 2002, 41 (7), 1892–1904. (13) Rogers, R. D.; Willauer, H. D.; Griffin, S. T.; Huddleston, J. G. J. Chromatogr., B: Biomed. Appl. 1998, 711 (1-2), 255–263. (14) Willauer, H. D.; Huddleston, J. G.; Griffin, S. T.; Rogers, R. D. Sep. Sci. Technol. 1999, 34 (6-7), 1069–1090. (15) Katritzky, A. R.; Ta¨mm, K.; Kuanar, M.; Fara, D. C.; Oliferenko, A.; Oliferenko, P.; Huddleston, J. G.; Rogers, R. D. J. Chem. Inf. Comput. Sci. 2004, 44 (1), 136–142.
Madeira et al. (16) Kamlet, M. J.; Taft, R. W. J. Am. Chem. Soc. 1976, 98 (2), 377– 383. (17) Taft, R. W.; Kamlet, M. J. J. Am. Chem. Soc. 1976, 98 (10), 2886– 2894. (18) Kamlet, M. J.; Abboud, J. L.; Taft, R. W. J. Am. Chem. Soc. 1977, 99 (18), 6027–6038. (19) Kamlet, M. J.; Abboud, J. L. M.; Abraham, M. H.; Taft, R. W. J. Org. Chem. 1983, 48 (17), 2877–2887. (20) Spange, S.; Keutel, D. Liebigs Ann. Chem. 1993, 981–985. (21) Marcus, Y. J. Chem. Soc., Perkin Trans. 2 1994, (8), 1751–1758. (22) Labban, A. S.; Marcus, Y. J. Chem. Soc., Faraday Trans. 1997, 93 (1), 77–79. (23) Trisˇovic´, N.; Banjac, N.; Valentic´, N.; Usˇc´umlic´, G. J. Solution Chem. 2009, 38 (2), 199–208. (24) Kim, I. W.; Jang, M. D.; Ryu, Y. K.; Cho, E. H.; Lee, Y. K.; Park, J. H. Anal. Sci. 2002, 18 (12), 1357–1360. (25) Singh, P.; Pandey, S. Green Chem. 2007, 9 (3), 254–261. (26) Huddleston, J. G.; Willauer, H. D.; Rogers, R. D. Phys. Chem. Chem. Phys. 2002, 4 (16), 4065–4070. (27) Madeira, P. P.; Teixeira, J. A.; Macedo, E. A.; Mikheeva, L. M.; Zaslavsky, B. Y. J. Chromatogr., A 2008, 1185 (1), 85–92. (28) Madeira, P. P.; Teixeira, J. A.; Macedo, E. A.; Mikheeva, L. M.; Zaslavsky, B. Y. Fluid Phase Equilib. 2008, 267 (2), 150–157. (29) Marcus, Y. Chem. Soc. ReV. 1993, 22 (6), 409–416. (30) Nicolet, P.; Laurence, C. J. Chem. Soc., Perkin Trans. 2 1986, (7), 1071–1079. (31) Marcus, Y. J. Phys. Chem. 1991, 95 (22), 8886–8891. (32) Zaslavsky, B. Yu.; Gulaeva, N. D.; Rogozhin, S. V.; Gasanov, A. A.; Masimov, A. A. Mol. Cell. Biochem. 1985, 65 (2), 125–130. (33) Rodriguez, O.; Silve´rio, S. C.; Madeira, P. P.; Teixeira, J. A.; Macedo, E. A. Ind. Eng. Chem. Res. 2007, 46 (24), 8199–8204. (34) Zaslavsky, B. Y.; Miheeva, L. M.; Gasanova, G. Z.; Mahmudov, A. U. J. Chromatogr., A 1987, 403 (C), 123–130. (35) Tanaka, H.; Kuboi, R.; Komasawa, I. J. Chem. Eng. Jpn. 1991, 24 (5), 661–664. (36) Kuboi, R.; Hasegawa, T.; Yano, K.; Komasawa, I. J. Chem. Eng. Jpn. 1995, 28 (6), 797–802. (37) Zaslavsky, B. Y.; Miheeva, L. M.; Rogozhin, S. V. J. Chromatogr., A 1981, 212 (1), 13–22.
JP907346S
