Thermal stability and enzymatic activity of RNase A in the presence of cationic gemini surfactants

International Journal of Biological Macromolecules 50 (2012) 1151–1157

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Thermal stability and enzymatic activity of RNase A in the presence of cationic gemini surfactants Razieh Amiri a , Abdol-Khalegh Bordbar a,∗ , Douglas V. Laurents b , Ahmad Reza Khosropour a , Iraj Mohammadpoor-Baltork a a b

Department of Chemistry University of Isfahan, Isfahan 81746-73441, Iran Instituto de Química Física Rocasolano, CSIC, Serrano 119, Madrid 28006, Spain

a r t i c l e

i n f o

Article history: Received 20 December 2011 Received in revised form 17 January 2012 Accepted 17 January 2012 Available online 24 January 2012 Keywords: Thermal denaturation Protein stability Enzymatic activity Ribonuclease A Gemini surfactant

a b s t r a c t The thermal stability and enzymatic activity of bovine pancreatic ribonuclease A (RNase A) have been investigated in the presence of a homologous series of cationic gemini surfactants (alkanediyl-␣,␻bis(hydroxyethyl methyl hexadecyl ammonium bromide)). UV, circular dichorism and fluorescence spectroscopies have been used for this study. The denaturation curves at various surfactant concentrations were analyzed on basis of a two-transition model to obtain values of Tm (temperature at the midpoint of denaturation) and Hm (enthalpy change at Tm ) of each transition. The main conclusion of this study is that these cationic gemini surfactants slightly activate and stabilize RNase A below their critical micelle concentrations at pH 5.0. The cationic gemini surfactant with the shorter spacer interacts more efficiently with RNase A than those with longer spacers. © 2012 Elsevier B.V. All rights reserved.

1. Introduction For many medical and biotech applications, it is desirable to increase protein stability and activity, which are the result of a balance between the intramolecular interactions of protein functional groups and their interaction with the solvent environment [1–3]. The practical use of modified protein enzymes with enhanced thermal stability for performing various tasks in cutting edge biotechnology is very promising [4–6]. For 35 years, the prevailing view has been that the hydrophobic effect is the dominant force in protein folding. Recently, the importance of hydrogen bonding has become clear. Studies of mutant proteins have improved our understanding of the forces stabilizing proteins [7–11]. Overall, the conformational stability of a protein is defined as the free energy change, G, for the equilibrium between the native state and the denatured state ensemble under physiological conditions [12,13]. The interaction between surfactants and different enzymes has been studied by using various techniques such as, electrochemistry, fluorescence, absorbance spectroscopy, conductometry, dynamic light scattering, resonance light scattering, circular dichroism and turbidity [14–18]. There are many extensive reports on the synthesis and investigation of varied properties of gemini

∗ Corresponding author. Tel.: +98 311 793271; fax: +98 311 6689732. E-mail addresses: [email protected], khalegh [email protected] (A.-K. Bordbar). 0141-8130/$ – see front matter © 2012 Elsevier B.V. All rights reserved. doi:10.1016/j.ijbiomac.2012.01.027

surfactants with different techniques [19–24]. These surfactants possess many remarkably qualities which could make them advantageous as membrane mimetics for biophysical studies, including low ␮M critical micelle concentrations (CMCs), slow millisecond monomer ↔ micelle kinetics and smaller micelle sizes [25]. Conventional surfactants, such as cetyltrimethylammonium bromide (CTAB) [26] or sodium dodecyl sulfate (SDS) denature most globular proteins and cause them to adopt a nonnative highly helical structure. This action is due to the interaction of the protein with SDS monomers, not SDS micelles [27]. Given the much lower CMCs of gemini surfactants, the concentration of their monomers is much lower and their effect on protein structure and function may well be different than that of SDS. Moreover, in recent years, gemini surfactants have begun to be widely used in biotech applications, which further increases interest in studies of their effects on protein structure and stability. Recently, the interaction of bovine serum albumin (BSA) with gemini surfactants was investigated and physicochemical properties of BSA were studied as a function of the spacer length [28–30]. Since BSA has an unusual ability to bind to many different hydrophobic ligands, it is interesting to also study the interaction of gemini surfactants, with other proteins with more typical properties. In contrast to BSA, bovine pancreatic ribonuclease A (RNase A) has a much lower tendency to bind nonpolar ligands. RNase A has an excess of cationic residues, especially Lys and is positively charged at neutral pH. This excess of cationic groups plays key roles in

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C2H4OH CH3

+

N

(CH2)S

C16H33

C2H4OH +

N

CH3 , 2Br

-

C16H33

Scheme 1. The chemical structure of alkanediyl-␣,␻-bis(hydroxyethylmethylhexadecyl ammonium bromide).

the binding of enzyme to its polyanionic substrate, single stranded RNA. This enzyme has played a crucial role as a model system in studies of protein structure, folding and unfolding pathways and enzyme catalysis [31,32]. This protein consists of 124 amino acid residues with a molecular mass of 13.7 kDa. In both the crystal and solution states, the protein consists of three ␣-helices and six ␤strands and overall, it contains about 23% ␣-helices, 46% ␤-sheets, 21% turns and 10% unordered structure [33]. Since the effects of pH, temperature, various salts and denaturants on the conformational stability of RNase A have been thoroughly studied [34,35]; this protein represents an excellent model system for studying the effects of gemini surfactants on protein conformational stability. In addition, the interaction between RNase A and monomeric surfactants such as the anionic surfactant SDS and the cationic surfactant, n-dodecyltrimethyl ammonium bromide (DTAB) has been investigated by absorption titration spectroscopy, differential scanning calorimetry and equilibrium dialysis experiments [26,27]. Also, the anionic surfactant sodium di-(2ethylhexyl) sulfosuccinate (AOT) has been used to purify RNase A [36]. However, until now no report has been published on the effect of cationic gemini surfactants on the overall structure and conformational stability of RNase A. In this work, we used a combination of fluorescence spectroscopy and absorption titration spectroscopy to determine the effect of a homologous series of alkanediyl-␣,␻-bis(hydroxyethyl methyl hexadecyl ammonium bromide) as cationic gemini surfactants on the native structure and thermal stability of RNase A. The thermal denaturation curves of RNase A in the presence of various amounts of these geminis were determined and analyzed in order to obtain the thermodynamic stability parameters of RNase A. We observed that these cationic gemini surfactants with CMC values 1.39 [37], 2.51 [38] and 2.40 ␮M [37] for butandiyl-1,4, pentanediyl-1,5-, and hexanediyl-1,6-, bis(hydroxyethyl methyl hexadecyl ammonium bromide), respectively, increases slightly the thermal stability and enzyme activity of RNase A at concentrations below their critical micelle concentration (CMC) at pH 5.0. The results also show that interaction of RNase A is most effective for the gemini surfactant with the shortest spacer. 2. Materials and methods 2.1. Materials 2(Methylamino) ethanol and n-hexadecyl bromide were purchased from Aldrich. 1,4-dibromo butane, 1,5-dibromo pentane and 1,6-diboromo hexane were bought from Merck. Cationic gemini surfactants alkanediyl-␣,␻-bis(hydroxylethyl methyl hexadecyl ammonium bromide) were synthesized by the method used by Devi and coworkers [37]. The general structure of these surfactants is shown in Scheme 1. Where s = 4, 5 and 6. In this work, we use the abbreviation GSX for this set of cationic gemini surfactants, and GSB, GSP and GSH for butanediyl-1,4-bis(hydroxyethyl methyl hexadecyl ammonium bromide), pentanediyl-1,5-bis(hydroxyethyl methyl hexadecyl ammonium bromide) and hexanediyl-1,6-bis(hydroxyethyl methyl hexadecyl ammonium bromide), respectively.

RNase A (EC 3.1.27.5) and cytidine 2 ,3 -cyclic monophosphate monosodium salt were purchased from Sigma. The buffer used was solution of 0.05 M citrate containing 0.1 M KCl at pH 5.0. The concentration of RNase A was determined spectroscopically using the molar absorption coefficient (M−1 cm−1 ) value of 9800 at 277.5 nm [39]. Cationic gemini surfactants were dissolved in Milli Q water and filtered through a 0.2 ␮m Millipore filter paper. Buffer solution was prepared with Milli Q water and filtered. 2.2. Fluorescence experiments The RNase A samples were prepared from stock solution by dissolving 1 mg of the enzyme into 1 mL of citrate buffer pH 5.0. The fluorescence experiments were done at room temperature, 23 ◦ C with a Shimadzu RF-5000 spectrofluorimeter. Both the excitation and emission slit widths were fixed at 5 nm. The excitation wavelength was 280 nm to excite the tyrosine residues. The initial concentration of RNase A in all solutions was 14.6 ␮M. All the fluorescence experiments were repeated at least three times. 2.3. Circular dichroism spectroscopy Far UV circular dichroism (CD) spectra at 200–260 nm intervals were done for RNase A solution (1 mg/mL) in the absence and presence of GSX in citrate buffer 0.05 M/0.1 M KCl pH 5.0, and at 25 ◦ C. These measurements were made using a 0.1 cm pathlength cuvette in a Jasco 810 CD spectropolarimeter in equipped with a Peltier module for computer controlled temperature adjustment. The scan speed and band width were 50 nm min−1 and 1 nm, respectively. The response time of these experiments was 4 s. 2.4. UV experiments 2.4.1. Absorbance titration spectroscopy First, a 1 mg/mL RNase A buffered solution was prepared and its absorption spectrum was obtained at room temperature. Thereafter, titrations were performed by addition of gemini surfactants to the solution containing RNase A. After each addition of gemini surfactants, the solution was incubated 90 s to permit it to reach equilibrium, prior to the measurement of the next UV spectrum. Absorbencies at max were corrected for dilution. All these experiments were repeated at least three times. 2.4.2. Thermal denaturation of RNase A Thermal denaturation studies were carried out with a Cary 500 UV–Vis instrument. Each sample was heated from 20 to 85 ◦ C at a rate of 1.0 ◦ C/min. Changes in absorbance with increasing temperature were followed at 287 nm. After thermal denaturation, each sample was recooled quickly to 20 ◦ C and the absorbance was remeasured. The initial and final absorbance values at 20 ◦ C were used to assess the reversibility of thermal denaturation. Each heatinduced transition curve was analyzed for Tm (temperature at the midpoint of denaturation) and Hm (enthalpy change at Tm ) using a non-linear least-squares analysis according to the equation Y (T ) =

(YN (T ) + YD (T )exp[−Hm /R(1/T − 1/Tm )]) 1 + exp[Hm /R(1/T − 1/Tm )]

(1)

where Y(T) is the experimentally observed absorbance of the protein at temperature T(K), YN (T) and YD (T) are the absorbance of the native and denatured molecules at T(K), R is the gas constant, Hm is the van’t Hoff enthalpy change at Tm . In the analysis of the transition curve, it was assumed that a linear function describes the dependence of the absorbance properties of the native and denatured protein molecules (YN = aN + bN T, YD = aD + bD T), where aN , bN , aD and bD are temperature independent coefficients [40,41]. Hm

R. Amiri et al. / International Journal of Biological Macromolecules 50 (2012) 1151–1157

and Tm were obtained from fitting transition curve points with Eq. (1). The fitting was done using Sigma Plot software, version 8.

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2

The enzymatic activity of RNase A was measured by the procedure described previously [42], with some modifications. Briefly, a 4.0 mg/mL stock solution of cytidine-2 ,3 -cyclic mono phosphate, was prepared in 0.05 M citrate buffer, pH 5.0. Then 650 ␮L of buffer solution was pipetted into each of the two 1 mL cuvettes. Then, 50 ␮L of enzyme solution (1.0 mg/mL) was added to both cuvettes, and the spectrophotometer reading was set to 0 at 288 nm. Next, 35 ␮L of buffer and substrate solutions were added to the reference and sample cuvettes, respectively. The content was mixed by inverting the cuvettes several times and the change in the absorbance at 288 nm was recorded over time. The difference in the absorbance for the initial rate was determined by extrapolation to time zero. The moles of substrate converted to product (mol S → P) per unit enzyme per unit time, was expressed as activity units. This was estimated from the kinetic data by dividing the initial rate by the RNase A concentration and multiplying by a constant as follows: U = 1 unit activity of RNase A

=

C



[RNaseA](mg mL−1 )

0

1

2

3

4

5

[GSX] (μM) Fig. 1. The variation of the relative fluorescence quantum yield of RNase A solution (14.6 ␮M) in the presence of various concentrations of GSB (), GSP (䊉) and GSH (). Q0 and Q are spectra integration of RNase A alone and in the presence of GSX, respectively. The measurements were done in 0.05 M citrate/0.1 M KCl buffer pH 5.0 and at 25 ◦ C. The excitation wavelength was 280 nm and band width was 5 nm for both excitation and emission. The data shown are one representative experiment of the three independent experiments that were performed.

3.3. UV measurements

OD288 t

 (2)

where C is constant for all of the measurements [43]. The enzyme activity in the presence of various amounts of GSX was measured as described above with the difference that both cuvettes contained an equal amount of GSX. The results were presented as relative activity of enzyme in the presence (U) and absence (U0 ) of GSX (U/U0 ). The assay was carried out at 25 ◦ C with a Shimadzu UV-160 spectrophotometer using 1 cm quartz cells. 3. Results 3.1. Fluorescence measurements Fig. 1 shows the variations of the fluorescence quantum yield intensity, Q/Q0 , of RNase A versus concentrations of GSB, GSP and GSH. A small increase in the fluorescence quantum yield intensity of RNase A without any shift in the emission wavelengths is seen upon addition of GSB, but no significant change is seen for GSP and GSH. This observation can be related to the more positive charge density of GSB that is due to close amine groups in it. This structural property causes GSB to be favor to bind with a pair of suitably located negatively charged groups of RNase A; that lead to more changes in microenvironment of Tyr residues and their quantum efficiency. 3.2. CD spectroscopy of RNase A in the presence of GSX Far UV-CD spectra were recorded at 25 ◦ C, pH 5.0 for RNase A alone and in the presence of GSX. Changes in CD at 222 nm, which are diagnostic for helical structure in RNase A [44] was followed by increasing of GSX. No definitive spectral evidence for protein/surfactant interactions was observed in this experiment under these conditions. The CD spectra of RNase A alone and with GSP are shown in Fig. 2. The similar results (not shown) were observed for GSB and GSH

UV difference spectra of RNase A at 287 nm in the presence of various concentrations of cationic gemini surfactants are plotted in Fig. 3 as variation in ε versus concentration of GSX. ε is defined as ε[GSX] − ε0 where ε[GSX] and ε0 , are the extinction coefficients of RNase A at a specified and zero concentrations of GSX, respectively. A linear increment in ε is observed upon increasing GSX concentrations. 3.4. Thermal denaturation measurements The heat-induced denaturation of RNase A in the presence of various concentrations of GSB, GSP and GSH was followed by observing changes in ε287 (Figs. 4 and 5). The transition curves for RNase A in the absence of GSX are characteristic of two state folded ↔ denatured transitions. Very similar results have been obtained in previous studies; see for example [45]. On the basis of the absorbance values measured at 20 ◦ C before and after heating, the thermal denaturation of RNase A, alone or in the presence of GSX, was found to be reversible.

0

-2000 -1



0

-4000

2

(mol S → P) (mol RNase A × t)

1

GSB GSP GSH

[Θ] deg.cm .dmol

=

Q/Q0

2.5. Enzymatic activity measurements

-6000

-8000 RNase A RNase A+GSP

-10000 200

210

220

230

240

250

260

Wavelength (nm) Fig. 2. The variation of CD spectrum of RNase A alone (dark line) and in the presence of cationic gemini surfactant GSP 0.94 ␮M (dashed line) in 0.05 M citrate/0.1 M KCl buffer pH 5.0 and at 25 ◦ C. The response time of these experiments was 4 s.

R. Amiri et al. / International Journal of Biological Macromolecules 50 (2012) 1151–1157

4

1

3

0

a

-1

Δε*10 /(M .cm )

1154

-1

-1

2

3

-2

1

RNase A RNase A + 0.37 μM GSB RNase A + 0.63 μM GSB RNase A + 0.88 μM GSB

-3 0

GSB GSP GSH

-4

-1 0

1

2

3

4

1

5

b

[GSX] (μM)

Interestingly, the transition shows an important difference in the presence of GSX. At low to intermediate temperatures (around 55 ◦ C), the ε287 first increases, but then decreases with further heating. This behavior is seen for all the surfactant molecules studied here. These data suggest that the thermal unfolding mechanism may be altered in the presence of these gemini surfactants so that the native protein passes through an intermediate state, with a high ε287 at 55 ◦ C before it completely unfolds at high temperatures. Therefore, the unfolding process would involve two distinct transitions: N ↔ I ↔ D. The concept that native state RNase A has two transitions in its unfolding profile has been demonstrated previously by Stelea et al. [46]. Those results ascribed the first transition to the unfolding of part of the secondary structure and the second transition to the loss of tertiary structure and the remainder of the secondary structure. The first transition, transition I, is centered in the temperature range of 20–55 ◦ C and is represented here by the reaction N ↔ I, where I is the thermodynamically stable intermediate state of the protein between its N (native) and D (denatured) states. The second transition, transition II, occurs in the temperature range of 50–85 ◦ C

0

Δε287×10-3 (M -1.cm-1 )

Fig. 3. Difference of RNase A extinction coefficients in the presence and absence of various concentrations of GSX (ε = ε[GSX] − ε0 ) at 287 nm. Extinction coefficients were corrected at max for dilution. The concentration of RNase A was 73 ␮M and all of the measurements were performed 0.05 M citrate/0.1 M KCl buffer pH 5.0 and at 25 ◦ C. The data shown are one representative experiment of the three independent experiments that were performed.

-1

-2

RNAse A RNAse A + 0.26 μM GSP RNAse A + 0.51 μM GSP RNAse A + 0.76 μM GSP

-3

-4 1

c

0

-1

-2

RNAse A RNAse A + 0.31 μM GSH RNAse A + 0.62 μM GSH RNAse A + 1.80 μM GSH

-3

-4 20

1

30

40

50

60

70

80

90

Temperature (°C)

0.5

Fig. 5. Representative thermal denaturation curves for RNase A alone (䊉) and in the presence of various sub-CMC concentrations of GSB (a), GSP (b) and GSH (c) which are given in the figures. The concentration of RNase A was 73 ␮M and the temperature scan rate was 1 ◦ C/min. The buffer used was 0.05 M citrate/0.1 M KCl pH 5.0. In order to maintain clarity all curves transitions are not shown.

0

-0.5

-1

-1.5 20

30

40

50

60

70

80

Temperature (°C) Fig. 4. Thermal denaturation profile of RNase A solution (73 ␮M) in the presence of GSP (1.8 ␮M) which is followed by ε287 (ε287 (T) − ε287 (20 ◦ C)) in 0.05 M citrate/0.1 M KCl buffer pH 5.0. The temperature scan rate was 1 ◦ C/min. Three dark lines represent two distinct transitions which the extrapolated baselines assuming that the temperature-dependence of the absorbance property is linear for each transition.

and is represented by the reaction I ↔ D. Assuming that each of these two processes N ↔ I and I ↔ D follows a two-state mechanism, the thermodynamics parameters, HI (the enthalpy change associated with the transition I), HII (the enthalpy change associated with the transition II), TmI (the temperature at the midpoint of first transition), TmII (the temperature at the midpoint of second transition) were determined and are given in Table 1. HmII is the difference between the HmII of RNase A in the presence and absence of GSX and similarly TmII is defined as the difference between the TmII of RNase A in the presence and absence of GSX.

R. Amiri et al. / International Journal of Biological Macromolecules 50 (2012) 1151–1157

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Table 1 Thermodynamic parameters for thermal denaturation of RNase A in the presence of GSX at pH 5.0. Concentration (␮M)

HmI (kJ mol−1 )

HmII (kJ mol−1 ) 449 ± 12

GSX

0.00

GSB

0.39

419 ± 44

GSB GSB GSB GSB GSB GSB

0.63 0.88 1.12 1.35 2.99 8.26

259 521 505 446 315 228

± ± ± ± ± ±

30 70 134 74 45 26

433 528 392 473 471 596

± ± ± ± ± ±

GSP GSP GSP GSP GSP GSP GSP

0.26 0.52 0.76 1.70 1.92 5.18 8.42

534 529 468 438 265 385 538

± ± ± ± ± ± ±

46 93 44 35 15 16 18

575 571 486 504 563 579 591

GSH GSH GSH GSH GSH GSH GSH GSH GSH

0.31 0.62 0.92 1.51 1.79 2.07 2.35 2.62 6.10

327 572 388 457 304 330

± ± ± ± ± ±

33 36 84 57 14 22

403 459 518 420 519 509 401 390 553

496 ± 7

HmII (kJ mol−1 )

TmI (◦ C)

TmII (◦ C)

0.00

TmII

65.19 ± 0.067

0.00

66.30 ± 0.031

1.11

−73.64

36.00 ± 0.23

15 14 8 12 9 18

−15.72 79.41 −56.34 23.89 22.14 147.20

40.18 37.83 37.40 37.27 42.78 35.63

± ± ± ± ± ±

0.42 0.23 0.50 0.34 0.56 0.62

66.49 66.85 66.21 65.77 66.01 67.04

± ± ± ± ± ±

0.084 0.054 0.057 0.057 0.042 0.055

1.30 1.66 1.02 0.58 0.83 1.86

± ± ± ± ± ± ±

19 16 12 6 12 9 12

126.57 122.66 36.93 55.49 114.05 126.68 142.13

35.86 34.53 35.02 36.05 36.18 36.21 34.64

± ± ± ± ± ± ±

0.15 0.29 0.18 0.17 0.20 0.10 0.06

65.78 66.04 66.30 66.60 66.44 66.86 67.00

± ± ± ± ± ± ±

0.052 0.272 0.062 0.026 0.040 0.032 0.038

0.59 0.85 1.12 1.41 1.25 1.67 1.82

± ± ± ± ± ± ± ± ±

6 8 10 5 7 7 36 40 16

−45.39 9.82 68.92 −28.75 70.14 60.05 −47.72 −58.44 104.20

33.03 37.49 34.65 37.86 38.55 38.54

± ± ± ± ± ±

0.27 0.10 0.53 0.25 0.14 0.18

66.13 66.47 66.98 67.07 66.72 66.31 62.81 65.53 66.36

± ± ± ± ± ± ± ± ±

0.04 0.04 0.04 0.03 0.03 0.03 0.24 0.29 0.06

0.94 1.28 1.79 1.88 1.53 1.12 −2.38 0.34 1.17

Table 2 RNase A relative activity in the presence of various concentrations of GSX at pH 5.0 and 25 ◦ C.a GSB

GSP

GSH

1.15 (0.26 ␮M) 1.17 (0.63 ␮M) 1.33 (0.88 ␮M) 1.06 (3.00 ␮M)

0.95 (0.37 ␮M) 1.03 (0.51 ␮M) 1.20 (0.76 ␮M) 0.98 (3.00 ␮M)

1.13 (0.31 ␮M) 1.33 (0.62 ␮M) 1.09 (1.80 ␮M) 1.04 (3.00 ␮M)

a In each column, the relative activity to RNase A alone, which is equal to 1.00, is given and the concentration of GSX is written within parentheses.

3.5. Enzymatic activity In Table 2 RNase A activity in the presence and absence of GSX at citrate buffer pH 5.0 has been compared. The activity of this enzyme rises in the presence of different concentrations of GSX below CMC. 4. Discussion We have used ultraviolet absorbance, fluorescence, CD spectroscopies and enzymatic activity measurements to determine the effect of cationic gemini surfactants on the structure, thermal stability and catalytic activity of RNase A. The addition of gemini surfactants produces small changes in the UV absorbance and null to slight alterations in the fluorescence emission of RNase A. The changes in UV absorbance and fluorescence emission are somewhat larger for GSB as compared to GSP or GSH, but in all cases are much smaller in magnitude than the changes expected for complete denaturation. Absorbance titration spectroscopy of RNase A in the presence of GSX shown in Fig. 3 indicates a very slight increase in ε with increase the concentration of GSX. This perturbation is stronger in the case of GSB, and we might suggest that this may arise from its two charged amine groups being held closer together in this surfactant, which could favor binding to a pair of suitably positioned negatively charged carboxylate groups such as Glu 2 & Glu 9 or Glu 49 & Asp 53. Based on the 3D structure of RNase A (Fig. 6), the binding of the quaternary ammonium groups of GSX to Glu 2 & Glu 9 or

Fig. 6. The 3D structure of ribonuclease showing Glu side chains in orange, Asp side chains in red, Tyr side chains in green, the backbone in white and backbone hydrogen bonds in brown. Glu 2 and Glu 9 (yellow balls) and Glu 49 and Asp 53 (blue balls) are positioned to bind the two quaternary amine groups of GSX while allowing the long hydrophobic surfactant tails to reach and possibly pack against Tyr 76, 92 and 115, which are partially exposed to solvent. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)

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Glu 49 & Asp 53 would be compatible with the aliphatic hydrocarbon gemini tails contacting hydrophobic moieties along the surface of native RNase A, including Tyr 76, Tyr 92 or Tyr 115, or some combination of these partially exposed Tyr, and not Tyr 25, 73 and 97, which are essentially buried [47]. Whereas its proof awaits future corroboration, this speculative hypothetical working model for the binding of sub-CMC concentrations of GSX to RNase A accounts for the data we report and is consistent with the body of knowledge on RNase A structure. Since the complete unfolding of RNase A is accompanied by ε287 changes of over 1000 cm−1 M−1 [48,49]; the much smaller changes observed here are evidence that GSX does not unfold RNase A at ambient temperatures. No significant change was observed in CD spectrum of RNase A in the presence of GSP 0.94 ␮M in Fig. 2. As regarding the CMC value of GSP is 2.51 ␮M, we can conclude that these cationic gemini surfactants GSX do not change secondary structure of RNase A at pH 5.0 in sub-CMC. Thus, whereas the simple anionic surfactants SDS can provoke the complete loss of native structure of RNase A, the results reported here are consistent with these cationic gemini surfactants binding to the surface of folded RNase A without provoking denaturation. The thermal denaturation studies reveal that these gemini surfactants affect the conformational stability of RNase A. Low surfactants concentrations (below CMC) produce a small increase in stability of RNase A against thermal denaturation. All thermodynamic quantities, given in Table 1, were obtained from the analysis of heat denaturation curves of RNase A in the presence and absence of various GSX concentrations at pH 5.0. The heat-induced denaturation involves two independent steps. The second step (I ↔ D) represents the main unfolding transition. Analysis of each transitions I and II according to Eq. (1) assumes that the transition between the native and intermediate states and transition between the intermediate and denatured states are twostate processes. Comparison of Tm and Hm of RNase A in the absence of GSX gives excellent agreement with the results of F. Ahmad and coworkers [45]. This agreement leads us to believe that our measurements of transition curves and their analysis for thermodynamic parameters are authentic and accurate. Thermal denaturation curves were monitored by ε287 , the difference molar absorption coefficient at 287 nm, which probes changes in the environment of tyrosine residues in RNase A. In order to check whether the two-state assumption holds in the presence of GSX in transitions I and II, thermal denaturation curves were also monitored by fDI and fDII , the normalized transition curves of thermal denaturation of RNase A in the presence of different concentrations of GSX. As shown in Fig. 5, TmII generally increases in the presence of GSX. This indicates that sub-CMC concentrations of these cationic gemini surfactants stabilize RNase A. The approximate CMC values for GSB, GSP and GSH are 1.39, 2.51 and 2.40 ␮M respectively, at 30 ◦ C. It can be seen in Table 1, that TmII values are positive at all cases except for GSH at 2.35 ␮M concentration. The inconsistency of this data with other 23 data points, most probably can be attributed to the experimental error. This does not effect in our general conclusion relevant to stabilization effect of gemini surfactants. However, no distinguished trend is observed for variation of TmII with increasing gemini surfactant concentration. This can be easily understandable considering that molar ratio of surfactant/protein is less than unity for most cases. Nevertheless, this does not alter our general conclusion corresponds to stabilization effect of gemini surfactant below its CMC. It has been previously shown that the preferential binding of compounds to folded proteins stabilizes them against denaturation [50–52]. Also, enzyme activity results showed that activity of RNase A increases in the presence of these monomeric cationic gemini surfactants (Table 2). Thus the stabilization and activation observed

here could arise from preferential binding of gemini surfactants to the native state of RNase A. Above the CMC, however, the TmII values begin to gradually decrease. Thus, whereas monomeric gemini surfactants stabilize RNase A, as the CMC is reached and micelles start to form, the inverse trend towards RNase A destabilization is observed. 5. Conclusions Ultraviolet absorbance and fluorescence spectroscopic changes of RNase A in the presence of GSX show GSB binds more readily to the surface of RNase A than GSP or GSH. Analyses of the heatinduced denaturation with the three state model and investigation of enzyme activity lead to the following conclusion; these cationic gemini surfactants stabilize and activate RNase A in the sub-CMC concentration range at pH 5.0. Acknowledgements Funding from the Research Council of the University of Isfahan and the Spanish Ministry of Science & Innovation (CTQ 2010-21567C02-02) are acknowledged. References [1] S.N. Timasheff, Annu. Rev. Biophys. Biomol. Struct. 22 (1993) 67–97. [2] P.H. Yancey, M.E. Clark, S.C. Hand, R.D. Bowlus, G.N. Somero, Science 217 (1982) 1212–1222. [3] Q. Zou, B.J. Bennion, V. Daggett, K.P. Murphy, J. Am. Chem. Soc. 124 (2002) 1192–1202. [4] M.W.W. Adams, R.M. Kelly, Chem. Eng. News 73 (1995) 32–42. [5] R. Jaenicke, G. Böhm, Curr. Opin. Struct. Biol. 8 (1998) 738–748. [6] R. Jaenicke, Eur. J. Biochem. 202 (1991) 715–728. [7] M.G. Coll, I.I. Protasevich, J. Torrent, M. Ribó, V.M. Lobachov, A.A. Makarov, M. Vilanova, Biochem. Biophys. Res. Commun. 265 (1999) 356–360. [8] E. Chatani, R.H. Functional, J. Biosci. Bioeng. 92 (2001) 98–107. [9] C.N. Pace, B.A. Shirley, M. McNutt, K. Gajiwala, FASEB J. 10 (1996) 75–83. [10] E.J. Rosario, G.G. Hammes, Biochemistry 8 (1969) 1884–1889. [11] N. Tanimizu, H. Ueno, R. Hayashi, J. Biosci. Bioeng. 94 (2002) 39–44. [12] C.N. Pace, Trends Biochem. Sci. 15 (1990) 14–17. [13] Z. Saadati, A.K. Bordbar, Protein J. 27 (2008) 455–460. [14] Z. Chen, G. Liu, M. Chen, Y. Peng, M. Wu, Anal. Biochem. 384 (2009) 337–342. [15] R.C. Lu, A.N. Cao, L.H. Lai, J.X. Xiao, Colloids Surf. B. 64 (2008) 98–103. [16] S.K. Mehta, K. Bhawna, K. Bhasin, A. Kumar, J. Colloid Interface Sci. 323 (2008) 426–434. [17] H. Xu, H.Y. Xiong, Q.X. Zeng, J. Li, W. Yan, S.F. Wang, Electrochem. Commun. 11 (2009) 286–289. [18] T. Yoshimura, K. Esumi, J. Colloid Interface Sci. 276 (2004) 231–238. [19] P.J.J.A. Buijnsters, C.L.G. Rodríguez, E.L. Willighagen, N.A.J.M. Sommerdijk, A. Kremer, P. Camilleri, M.C. Feiters, R.J.M. Nolte, B. Zwanenburg, Eur. J. Org. Chem. 8 (2002) 1397–1406. [20] M. Deng, J. Li, J. Liu, X. Ma, Y. Wang, Colloids Surf. A 356 (2010) 97–103. [21] K. Esumi, M. Miyazaki, T. Arai, Y. Koide, Colloids Surf. A 135 (1998) 117–122. [22] M.A.-E. Jouani, S. Szonyi, S.Y. Dieng, A. Cambon, S. Geribaldi, New J. Chem. 23 (1999) 557–562. [23] F. Li, M.J. Rosen, S.B. Sulthana, Langmuir 17 (2001) 1037–1042. [24] Ø. Rist, A. Rike, L. Ljones, P.H. Carlsen, J. Molecules 6 (2001) 979–987. [25] X. Cui, X. Yang, H. Chen, A. Liu, S. Mao, M. Liu, H. Yuan, P. Luo, Y. Du, J. Phys. Chem. 112 (2008) 2874–2879. [26] M.N. Jones, H.A. Skinner, E. Tippingt, A. Wilkinson, Biochem. J. 135 (1973) 231–236. [27] A.A. Moosavi-Movahedi, M. Gharanfoli, K. Nazari, M. Shamsipur, J. Chamani, B. Hemmateenejad, M. Alavi, A. Shokrollahi, M. Habibi-Rezaei, N.S.C. Sorenson, Colloids Surf. B 43 (2005) 150–157. [28] M.A. Mir, J.M. Khan, R.H. Khan, G.M. Rather, A.A. Dar, Colloids Surf. B 77 (2010) 54–59. [29] S. Tardioli, A. Bonincontro, C.L. Mesa, R. Muzzalupo, J. Colloid Interface Sci. 347 (2010) 96–101. [30] D. Wu, G. Xu, Y. Sun, H. Zhang, H. Mao, Y. Feng, Biomacromolecules 8 (2007) 708–712. [31] J.L. Neira, M. Rico, Fold. Des. 2 (1997) R1–R11. [32] R.T. Raines, Chem. Rev. 98 (1998) 1045–1065. [33] A. Wlodawer, N. Borkakoti, D.S. Moss, B. Howlin, Acta Crystallogr. B 42 (1986) 379–387. [34] F. Ahmad, C.C. Bigelow, J. Mol. Biol. 131 (1979) 607–617. [35] D. Constantinescu, H. Weingartner, C. Herrmann, Angew. Chem. Int. Ed. 46 (2007) 8887–8889.

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