Buffers more than buffering agent: introducing a new class of stabilizers for the protein BSA

1114 | Phys. Chem. Chem. Phys., 2015, 17, 1114--1133 This journal is© the Owner Societies 2015

Cite this:Phys.Chem.Chem.Phys.,

2015, 17, 1114

Buffers more than buffering agent: introducinga new class of stabilizers for the protein BSA†

Bhupender S. Gupta,a Mohamed Tahab and Ming-Jer Lee*a

In this study, we have analyzed the influence of four biological buffers on the thermal stability of bovine

serum albumin (BSA) using dynamic light scattering (DLS). The investigated buffers include 4-(2-

hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES), 4-(2-hydroxyethyl)-1-piperazine-propanesulfonic

acid (EPPS), 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid sodium salt (HEPES–Na), and 4-morpholine-

propanesulfonic acid sodium salt (MOPS–Na). These buffers behave as a potential stabilizer for the native

structure of BSA against thermal denaturation. The stabilization tendency follows the order of MOPS–Na 4

HEPES–Na 4 HEPES c EPPS. To obtain an insight into the role of hydration layers and peptide backbone

in the stabilization of BSA by these buffers, we have also explored the phase transition of a thermoresponsive

polymer, poly(N-isopropylacrylamide (PNIPAM)), a model compound for protein, in aqueous solutions of

HEPES, EPPS, HEPES–Na, and MOPS–Na buffers at different concentrations. It was found that the lower

critical solution temperatures (LCST) of PNIPAM in the aqueous buffer solutions substantially decrease

with increase in buffer concentration. The mechanism of interactions between these buffers and protein

BSA was probed by various techniques, including UV-visible, fluorescence, and FTIR. The results of this

series of studies reveal that the interactions are mainly governed by the influence of the buffers on the

hydration layers surrounding the protein. We have also explored the possible binding sites of BSA with

these buffers using a molecular docking technique. Moreover, the activities of an industrially important

enzyme a-chymotrypsin (a-CT) in 0.05 M, 0.5 M, and 1.0 M of HEPES, EPPS, HEPES–Na, and MOPS–Na

buffer solutions were analyzed at pH = 8.0 and T = 25 1C. Interestingly, the activities of a-CT were

found to be enhanced in the aqueous solutions of these investigated buffers. Based upon the Jones–

Dole viscosity parameters, the kosmotropic or chaotropic behaviors of the investigated buffers at 25 1C

have been examined.

Introduction

Proteins are complex macromolecule systems with substantialstructural variability in their folded states. The proper function-ing of proteins, particularly in aqueous solution, is highly depen-dent on the pH value of the solution. Any fluctuation from thesuitable pH range can completely change the functional proper-ties of proteins. Many life processes, involving enzymatic reac-tions, function only in a narrow range of pH values. Thus, buffersolutions are essential in pH-dependent chemical reactions tomaintain the required pH values of the reaction medium.Proteins in aqueous solution are never found without a buffer.To maintain a suitable pH range, buffer compounds ideally

release or absorb hydronium ions in the reaction medium,according to the requirement.

On the basis of certain criteria for a suitable biological buffer,such as good solubility in water, low ion effect, low absorbance oflight at visible or ultraviolet wavelength, good stability and highpurity, Good and co-workers1–5 proposed a different class of bio-logical buffers derived from N-substituted aminosulfonic acids.These buffers are commonly known as Good’s buffers and providegood coverage in the physiological pH range (6 to 10). Indeed, theyare widely used in many research areas, including biology,biochemistry, and environmental studies.24 In addition to theabovementioned criteria for a suitable biological buffer, anotherimportant aspect is that the buffer should be inert, i.e., it shouldnot interfere with the reaction system. However, besides main-taining a constant pH value, many of these buffers are found tobe reactive, and their interactions with many metal ions havebeen reported by several investigators.6–9

Stellwagen et al.10 studied the influence of some neutral pH,amine-based buffers, TAE (Tris-acetate–EDTA) and TBE (Tris-borate–EDTA), on the capillary electrophoresis movement of a

a Department of Chemical Engineering, National Taiwan University of Science and

Technology, 43 Keelung Road, Section 4, Taipei 106-07, Taiwan.

E-mail: [email protected]; Fax: +886-2-2737-6644; Tel: 886-2-2737-6626b CICECO, Departamento de Quımica, Universidade de Aveiro, 3810-193 Aveiro,

Portugal

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c4cp04663c

Received 14th October 2014,Accepted 5th November 2014

DOI: 10.1039/c4cp04663c

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DNA molecule in a mixture of plasmid-sized DNA molecule anda small DNA oligonucleotide. The behavior of the DNAs wasfound to be unusually different in the presence of the respectivebuffers, which was attributed to the different interactions of thebuffers with the DNA molecule. Kameoka et al.11 observed thedifferent aggregation behavior of humanized immunoglobulinG (IgG) protein with various buffer species under the samephysiological conditions. For example, in the presence of MESand MOPS buffers, the aggregation propensity of the humanizedantibody appeared to be lower than in the presence of phosphatecitrate buffers. Welinder et al.12 reported that MES buffer facili-tates the formation of uniform gold nanoparticles. Ursi et al.13

found the deleterious effect of organic amine-based TRIS bufferon the growth rates and pigment contents of agarophytesGracilaria birdiae (Plastino and Oliveira). Kejnovsky and Kypr14

mentioned that TRIS molecule protects DNA from nicking uponirradiation with ultraviolet light. However, the protective effectwas only effective for the DNA backbone (not for the bases) inaqueous solution (not in formamide solution). Lubas et al.15

studied the activity of endo-a-D-mannosidase enzyme in thepresence of various biological buffers, and found the enhancedactivity of enzyme with 4-morpholine-ethanesulfonic acid (MES)and 3-morpholino-2-hydroxypropanesulfonic acid (MOPSO) buffersat pH 7.0 over buffers, such as HEPES or EPPS, while a negativeeffect on the activity of the enzyme was observed usingtris(hydroxymethyl)aminomethane (TRIS) buffer. Kaushal andBarnes16 performed the assays of bovine serum albumin (BSA)protein in the presence of several zwitterionic buffers usingbicinchoninic acid (BCA) for detection. Among the testedzwitterionic buffers, only 3-(cyclohexylamino)-1-propanesulfonicacid (CAPS) was found to have enhanced the absorbance of BSA,and it was found that the other tested buffers, such as HEPES,EPPS, 2-[(2-hydroxy-1,1-bis(hydroxymethyl)ethyl)amino]ethane-sulfonic acid (TES), TRIS, N-[tris(hydroxyl-methyl)methyl]glycine(TRICINE), and N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonicacid (BES), reduced the absorbance, whereas MES and MOPSexhibited negligible effects on the absorbance of BSA in thebicinchoninic acid protein assay. Therefore, the selection of aproper buffer is a challenging task, and it always requiresextensive testing.4

Despite the plenty of research being carried out involvingproteins in buffer solutions, relatively little information hasbeen reported on the mechanism of the protein–buffer inter-actions. The adverse effect of such interactions on stabilizing ordestabilizing protein structure is still unclear. Ugwu and Apte17

reported that the interaction between buffers and protein caneither stabilize or destabilize proteins. Generally, if a certainbuffer preferentially binds to the native state of a protein, itwould enhance the protein stability. In contrast, if a bufferselectively binds to the denatured state, it would decrease thestability. In addition, alterations in microenvironment or watershell around the protein molecule by buffers may also cause achange in the conformation of protein molecule. Because buffersalways serve as a chief component in vitro, understanding theinteractions of proteins with buffers is important. The knowledgeof such interactions can provide information about the binding

and the naturation or re-naturation effect of buffers on proteinstructure, as well as on the stability of proteins.

Proteins are complex molecules and they remain functionallyactive only in their native globular conformation at a specific pHvalue and temperature. This functional stability of the native stateof proteins results from the different types of interactions, such ashydrophobic, hydrogen bonding, van der Waals, electrostatic, andlocal peptide–peptide interactions.18 All these interactions arehighly dependent on temperature; thus, with increase in tempera-ture, proteins start to deviate from their native conformationand lose their conformational stability and become biologicallyinactive. Recently, several therapeutic protein products havebeen developed due to the advancement in recombinant DNAtechnology. However, their applications are limited becauseof the sensitivity of proteins towards chemical or physicaldegradation.19–22 Indeed, protecting the proteins from degradationand improving their stabilities are most desirable, particularly inpharmaceutical industry.

Therefore, as a part of our research, we attempt to find newpossibilities for stabilizing a globular protein BSA by buffers, and toexplore the mechanism of the interactions between the buffers andproteins. In our previous studies,23–25 we reported the influence ofsome important biological buffer compounds on the stability ofproteins in aqueous solutions using various techniques. In thisstudy, we have analyzed the tendency of four commonly usedbiological buffers, namely, 4-(2-hydroxyethyl)piperazine-1-ethane-sulfonic acid (HEPES), 4-(2-hydroxyethyl)-1-piperazinepropane-sulfonic acid (EPPS), 4-(2-hydroxyethyl)piperazine-1-ethanesulfonicacid sodium salt (HEPES–Na), and 4-morpholinepropanesulfonicacid sodium salt (MOPS–Na), to protect the native structure of BSAagainst thermal denaturation. These selected biological buffers(HEPES, EPPS, and HEPES–Na) are structurally related compounds,and all contain a piperazine ring and belong to the piperazinefamily. The buffer MOPS–Na contains a morpholine ring, and thusbelongs to the morpholine family. These buffers are suitable for aphysiological pH range from 6.8 to 8.7, and are widely used invarious biological applications.

To examine the thermal stability of BSA in the presence ofrespective buffers, we measured the hydrodynamic diameter ofBSA in 0.05 M, 0.5 M, and 1.0 M of HEPES, EPPS, HEPES–Na,MOPS–Na buffer solution at pH = 7.0, in the temperature rangeof 25 1C–75 1C, using a dynamic light scattering (DLS) instru-ment. The results of the DLS measurement indicate that thecommon biological buffers can serve as a potential stabilizer forthe native structure of BSA molecule. To explore the mechanismof this stabilizing nature of BSA in aqueous solutions of theinvestigated buffers, we probed the interaction of the studiedbuffers with BSA by means of molecular docking and spectro-scopy techniques (UV-visible, fluorescence, and FTIR).

Because proteins are primarily composed of polypeptidechains, any change in the native structure of proteins due toexternal or internal stress is mainly governed by their peptidebackbone. Therefore, the investigation of the influence of bufferson the peptide backbone unit can be useful. This information iscrucial for clarifying the mechanism of the interaction of proteinswith biological buffers. For this purpose, we have selected a

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well-known polymer poly(N-isopropylacrylamide) (PNIPAM), andanalyzed its interaction with the investigated buffers. PNIPAM is awell-known thermoresponsive amphiphilic polymer, which isused worldwide as a model for proteins.27 It is an isomer ofpoly(leucine) and possesses a pendant peptide group in the sidechain, and not in the backbone. Due to this peculiar structuralarrangement of PNIPAM, the major accessible interaction sites forbuffers are the peptide groups in the side chains. Otherwise,no any other hydrogen bond donating- or accepting-groups areavailable in this polymer.

The aqueous solution of PNIPAM exhibits a lower criticalsolution temperature (LCST) of around 32–35 1C.26 At tempera-tures below the LCST value, the polymer exists in a random coilconformation. On heating this polymer above the LCST value,the network of inter-molecular hydrogen bonding between thepolymer and water molecules is disrupted, which leads to thephase transition of PNIPAM from a random coiled conforma-tion to a compact globular form. The formed globular structureis simultaneously converted to optically detectable reversibleaggregates. This fascinating property of PNIPAM is investigatedto a great extent around the world because of its certainsimilarity to many biological systems, such as protein folding,native DNA packing, and network collapsing.27 Therefore, toexplore the role of hydration layers and the influence of thesebuffers on the peptide moiety of PNIPAM, we examined theLCST of PNIPAM in 0.0, 0.3, 0.5, 0.7, and 0.9 M HEPES, EPPS,HEPES–Na, and MOPS–Na buffer solutions by means of DLSand FTIR spectroscopy techniques. Moreover, this polymer isacclaimed in industries for its applications in various fields,such as drug or dye delivery,28,29 tissue engineering,30 tem-plates for biosensing,31 nanoparticles,32 and catalysis.33 Severalstudies were performed to elucidate the effects of salts,34–37

surfactants,38–42 sugars,43 cosolvents,44–65 urea,66 and ionicliquids67 on the phase transition of PNIPAM in aqueous media.As a consequence, the information about the LCST of PNIPAM inthe presence of buffer can be extremely useful from an industrialpoint of view. To find more experimental evidence for theinteractions between water and the investigated buffers, weobtained the viscosity coefficients (A and B) of the Jones–Doleequation68 for each buffer–water system at 25 1C and interpretedthe results in terms of the kosmotropic (structure maker) orchaotropic (structure breaker) nature of the solute buffer for thesolvent water. The Jones–Dole equation is widely used todescribe solute–solute and solute–solvent interactions.69–72

We selected BSA as a model protein because of its numerouswell-known functional applications. It serves as the majortransport carrier for drugs and both endogenous and exogenoussubstances. BSA has been utilized in various biochemical reac-tions; it is suitably absorbed on the surface of other substances,and it is capable of binding with other molecules, thus formingmolecular aggregates.73 Globular proteins, particularly BSA, arewidely used in various technical applications.74–77 However, theproper functional efficiency of proteins depends upon severalfactors, such as molecular structure, chemical environment andthermal stability.78–81 BSA has also been used as a functionalintegral in various food and health care products. The molecular

weight of BSA is 66 kDa. A single polypeptide chain of BSAmolecule contains 583 amino acid residues.82 It has threehomologous domains (I, II, and III), 17 disulfide bridges andone free SH group, which divide the protein into 9 loops(L1–L9).83 The structure of BSA has two tryptophan (Trp),20 tyrosine (Tyr), and 27 phenylalanine (Phe) aromatic aminoacid residues. The presence of these aromatic amino acidresidues, especially Trp134 in the first domain and Trp 212 inthe second domain, make this protein suitable for the study oftheir interactions with different ligands.84 Due to these specialstructural features, BSA was considered as a model protein forthe in vitro investigation of protein–drug interaction.

CT is used as an important biological enzyme for under-standing the mechanism of protein folding or unfolding in thepresence of cosolvents.85 The molecular weight of CT is 25 kDa.Its polypeptide chain is composed of 245 amino acid residues.It is composed of two juxtaposed-barrel domains with catalyticresidue bridging and disulfide bridges combining the threepolypeptide chains. The enzymatic site, commonly known ascatalytic triad, which is found in the second domain of CT, isformed by three amino acid residues: His 57, Asp 102, and Ser195.86,87 The structure of CT includes five disulfide bonds: Twodisulfide bonds, Cys1–Cys122 and Cys42–Cys58, are situated closeto the catalytic center. One disulfide bond, Cys191–Cys220, issituated near the renowned surface binding site, which is locatedin the vicinity of the catalytic triad. The other two disulfide bonds,Cys136–Cys201 and Cys168–Cys182, are observed away from thecatalytic center, as well as in the surface binding pocket.

Materials and methodsMaterials

The protein bovine serum albumin (BSA/fraction V, pH = 7.0) wasobtained from Acros Organics (USA). PNIPAM (Mn = 20 000–25 000),a-chymotrypsin (a-CT) from bovine pancreas type II, essentially saltfree, and the buffers, HEPES, EPPS, HEPES–Na, and MOPS–Na,each with a mass fraction purity 40.99, were purchased fromSigma Chemical Co. (USA). All the materials were used asreceived, without further purification. Aqueous solutions ofbuffer were prepared by dissolving the buffer in double distilledde-ionized water, which was obtained from a Nano pure-Ultrapure water purifying system with a resistivity of 18.3 MO cm. Allthe analysis samples of PNIPAM or BSA were gravimetricallyprepared using an electronic balance (R & D, Model GR-200)with an uncertainty of �0.1 mg.

Dynamic light scattering (DLS)

The hydrodynamic diameter (dH) and the size distribution ofBSA or PNIPAM in HEPES, EPPS, HEPES–Na, and MOPS–Nabuffer solutions were measured using dynamic light scattering(DLS) (ZetasizerNano ZS90, Malvern Instruments Ltd., UK).This instrument was equipped with a thermostatic samplingchamber for maintaining a constant temperature within therange of 0 1C–75 1C. A He–Ne laser light with a voltage of 4 mWand at a fixed wavelength (l) of 633 nm was used as a light source.

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The DLS measurements were performed at a fixed scatteringangle of 12.81, 901 or 1731. The averaged dH and the sizedistribution of the analyzed samples from the scattering intensitydata were calculated by built-in Malvare computer software. Thesamples for the DLS measurements were prepared with 20 mg cm�3

of BSA in 0.05, 0.5, and 1.0 M HEPES, EPPS, HEPES–Na, andMOPS–Na buffer solutions at pH = 7. The samples of PNIPAMwere prepared with 6 mg cm�3 of PNIPAM in 0 M to 0.9 Minvestigated buffer solutions. The analysis samples were filteredby passing through a 0.22 mm disposable filter (Millipore); about1.5 cm�3 of the BSA–PNIPAM solution, free from bubbles, wasplaced into a square glass cuvette with a round aperture(PCS8501) using a syringe. To protect the sample from contami-nation, we closed the mouth of the sampling cell with a Teflon-coated screw cap. Before performing the measurements, theBSA–PNIPAM samples were incubated for 4 to 5 hours at 25 1C toattain complete equilibrium. At least three repeated measure-ments were made for each sample at a given temperature.

Calculation of the Jones–Dole parameters

The Jones–Dole equation is recognized as an efficient way toestimate solute–solute and solute–solvent interactions.69–72 Thisequation is given as follows:

Z=Z0ð Þ � 1½ �C1=2

¼ Aþ BC1=2 (1)

where Zo and Z are the viscosities of the solvent and solution,respectively. A and B are the Jones–Dole viscosity parameters,which are the characteristics of solute–solute and solute–solventinteractions, respectively. C represents the buffer concentration(mol litre�1) of the solution. The values of A and B for each buffer–water system at 25 1C were estimated by the least square fittingmethod.

To obtain the Jones–Dole equation parameters, the experimentaldata of densities and viscosities for the series of aqueous buffersolutions were measured at 25 1C. The aqueous samples of eachinvestigated buffer (0.0 M, 0.2 M, 0.4 M, 0.6 M, 0.8 M and 1.0 M)were gravimetrically prepared. Prior to the analysis, the analysissamples were filtered through a 0.22 mm disposable filter (Millipore,Millex-GS). Densities (r) of all the analysis samples were measuredat atmospheric pressure using a digital vibrating U-tube densimeter(DMA 4500, Anton Paar, Austria). The kinematic viscosities (n)were experimentally determined with a microviscometer (Lovis2000 M/ME, Anton Paar, Austria). This viscometer automaticallyprovides the value of the viscosity (Z) of the analysis sample basedon the input value of density using the following equation:

Z = n�r (2)

At least three repeated measurements were made for eachsample and the uncertainties for the density and viscositymeasurements were estimated to be about �5 � 10�5 g cm�3

and �0.05 mPa s, respectively.

Fourier transform infrared (FTIR) spectra

The FTIR spectra of the samples containing PNIPAM or BSA inthe buffer solutions were measured using a Bio-Rad Digilab

FTS-3500 spectrometer with a spectral resolution of 8 cm�1 atan average of 300 scans. The polymer samples were preparedwith 20 mg cm�3 of PNIPAM in 0.0 M, 0.1 M, 0.5 M, and 0.7 Mbuffer solutions, whereas the protein samples were preparedwith 30 mg cm�3 of BSA in 0.05 M, 0.5 M, and 1.0 M buffersolutions at pH = 7. Each sample was placed between two ZnSewindows, 32 mm in diameter (Sigma-Aldrich), and a spacer,0.015 mm in thickness (EZ copper). The IR cell was attached toa metal cell holder, provided with a temperature controller(Model HT-32 heated transmission cell). The analysis of themeasurement results was executed using Varian Resolutions(Version 4.10) software. Prior to the spectrum measurement ofeach sample, the spectrum of the background solvent wasrecorded to avoid the interference of the solvent.

UV-vis spectra

The UV-visible absorption spectra were recorded at roomtemperature for 3.5 mg cm�3 of BSA in 0.05 M, 0.5 M, and1.0 M buffer solutions (pH = 7.0) using a UV-visible spectro-photometer (JASCO, V-550) with a 1.0 cm quartz cuvette. Theoperating conditions during the measurement were as follows:scan speed 100 nm min�1, scan range 190–600 nm, slit width2 nm, and Dl = 0.1 nm.

Fluorescence spectra

The emission spectra were measured at room temperature for3.5 mg cm�3 of BSA in 0.05 M, 0.5 M, and 1.0 M buffer solutionsat pH = 7.0 using an RF-5301PC Spectro-fluorophotometer(SHIMADZU), with a 1.0 cm quartz cell. The spectra wererecorded at excitation wavelengths of 230 nm and 280 nm withan excitation and emission slit width of 5 nm for the emissionrange of 250 nm to 450 nm.

Molecular docking

To obtain the most probable interaction site for the investi-gated buffers in protein BSA, the molecular docking calculationwas performed with Molegro Virtual Docker v. 4.0.88,89 Thecrystal structure of BSA was obtained from a protein data bank(PDB 3v03).90 The structure of each buffer was optimized in a gasphase by a molecular mechanics method using the AMBER91

force field using a Gaussian 0992 suite of computer software.The crystalline structure of BSA, free from water molecules,and the optimized structure of buffer molecule in the docking-Wizard were assigned to the required properties, such as bonds,hybridization, explicit hydrogen atoms, charges, and flexibletorsions, if missing and checked to ensure zero error. Thedocking parameters, such as energy threshold, maximum iteration,maximum population size, and grid resolution, were set to 100,1500, 50, and 0.3 Å, respectively. RMSD threshold for multiplecluster poses was set to less than 1 Å. Using the inbuilt searchalgorithms, the maximum 18 potential binding sites (cavities)with different volumes were detected in the entire BSA molecule.For each buffer, 10 optimum binding modes (docking pose) wereobtained corresponding to each cavity, with a sphere sufficientlylarge to accommodate the cavity centered on the binding sitesand 50 independent runs. Therefore, for each buffer molecule we

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obtained 180 binding modes with the entire BSA molecule, and thenthe best binding pose of each buffer (HEPES, EPPS, HEPES–Na, andMOPS–Na) out of the obtained 180 poses with BSA was selected onthe basis of the highest ranking docking score.

Activity measurement

The catalytic activities of an enzyme a-chymotrypsin (a-CT) in0.05 M, 0.5 M, and 1.0 M aqueous buffer solutions of HEPES,EPPS, HEPES–Na, and MOPS–Na were measured by estimatingthe appearance of p-nitrophenoxide at 400 nm, at pH = 8 and at25 1C with the help of a UV-visible spectrophotometer (JASCO,V-550). The detailed method to measure the activities is describedin the literature.93,94 The samples for the activity measurementswere prepared with 2 mg cm�3 of a-CT in 0.05 M, 0.5 M, and1.0 M HEPES, EPPS, HEPES–Na, and MOPS–Na buffer solutionat pH = 8. Prior to the activity measurements, the preparedenzyme-buffer samples were incubated for about 3 to 4 hours at25 1C to attain the complete equilibrium. To minimize theinstantaneous aqueous hydrolysis of PNPA, the stock solutionof substrate PNPA (4 mM) was prepared in acetonitrile. Thisstock solution of PNPA was used to measure the activity ofenzyme a-CT in different concentrations of buffer. The unit ofactivities was expressed as millimole of the product formed perunit time per gram of protein. The value of the activity wasobtained from at least three repeated measurements for eachsample at a given temperature.

Results and discussionDynamic light scattering (DLS)

In the present study, the influence of the commonly usedbiological buffers (HEPES, EPPS, HEPES–Na and MOPS–Na)on the thermal denaturation of the protein BSA was analyzedwith DLS. DLS technique works on the basis of the Stokes–Einstein principle and determines the hydrodynamic diameter(dH) of the particles in the solution from the variation in thescattered intensity of light with time. Each protein is knownto possess a characteristic conformational and structuralarrangement. Thus, different proteins exhibit different stabilitybehavior. Various types of non-covalent interactions, such ashydrogen bonding, electrostatic, and hydrophobic interactions,account for the stability behavior of proteins and maintain thestructure in a globular form against any type of external orinternal impetus. When any protein is subjected to heat, thenetwork of these temperature-dependent interactions starts tobecome weak. As a result, at a particular temperature, proteinmolecules start to unfold; this is known as critical or denatura-tion temperature (TC). A successive unfolding in the protein,due to the continuous increase in temperature over the char-acteristic TC value of the protein, causes the interior hiddenhydrophobic residues on the different chains of the sameprotein to become available for interacting with each other.These nonspecific interactions result in the formation of irrever-sible aggregates.95–97 The unfolding or aggregate formation leadsto a change in the hydrodynamic diameter (dH) and scattering

intensity of the protein. A marked point corresponding to thesignificant increase in the size and scattering intensity wasconsidered as the denaturation point of protein, which can beeffectively observed by DLS. Thus, this technique is highlyresponsive for the folding and unfolding of proteins, and ithas been used worldwide in the field of protein folding.98–101 Inaddition, DLS can also provide information about the intensitydistribution of the molecules of different shapes and sizes inthe solution.102,103 Therefore, the variation of the hydrodynamicdiameter of BSA and its intensity distribution as a function oftemperature in 0.05 M, 0.5 M, and 1.0 M HEPES, EPPS, HEPES–Na, and MOPS–Na buffer solutions, at pH = 7.0, were measuredusing DLS in the present study. The results of the measurementsare graphically presented in Fig. 1, and the measured values areprovided in Tables S.1 to S.4 in the ESI.†

On the basis of several previous investigations,104,105 aboutthe heat induced denaturation of BSA, it can be revealed that theconformational change of BSA molecule in aqueous solution isreversible in the temperature range of 25 1C–50 1C. This provesthat BSA always exists in the native state below 50 1C. Theunfolding of BSA molecule starts at 55 1C. The formation ofirreversible aggregates occurs at temperatures above 55 1C.A gel-type solid appears due to the successive unfolding ofBSA at temperatures above 70 1C.

From the tabulated values of dH in Tables S.1 to S.4 (ESI†)and the melting curves of BSA in HEPES, EPPS, HEPES–Na, andMOPS–Na (Fig. 1), it can be seen that the TC value of the nativeBSA in 0.05 M HEPES, HEPES–Na, and MOPS–Na buffers is55 1C. This means that BSA in 0.05 M concentration of theseinvestigated buffers cannot exist in the folded globular state at55 1C. To confirm the irreversible folding nature, the sampleswere cooled from their respective denaturation temperature(TC) to 25 1C, and no any change in the size was observed. Thedenaturation of BSA in 0.5 M and 1.0 M HEPES buffer solutionoccurred at 58 1C and 59 1C, respectively. The denaturationtemperature of BSA in 0.5 M and 1.0 M HEPES–Na buffersolution was found to be 60 1C and 62 1C, respectively. Thedenaturation of BSA in 0.5 M and 1.0 M MOPS–Na was observedto be 63 1C and 65 1C, respectively. Thus, these results indicatethat HEPES, HEPES–Na, and MOPS–Na buffers are suitable toprotect the native structure of BSA against thermal denaturation.The thermal stability of BSA increased with increase in respectivebuffer concentration. The enhanced tendency of HEPES–Na forstabilizing BSA over HEPES can be attributed to the more polarnature of HEPES–Na than HEPES. The denaturation temperatureof BSA in 0.05 M or 0.5 M EPPS is 55 1C and it is changed to 56 1Cin 1.0 M EPPS buffer solution. It was unfortunate to see thatalthough EPPS buffer appeared to interact strongly with BSA(as it appeared from the UV-visible and florescence spectra, andthe molecular docking analysis), it failed to protect the nativestructure of BSA against thermal denaturation to a considerableextent. Nevertheless, the denaturation temperature of BSA in thehighest concentration of the investigated buffers was significantlyshifted to a higher temperature. The overall stabilizing tendencyof these investigated buffers for BSA in aqueous solution at pH = 7follows the order of MOPS–Na 4 HEPES–Na 4 HEPES c EPPS.

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These results clearly indicate the ability of these investigatedbuffers for stabilizing the protein BSA against thermal denaturation.

In our previous investigations,23–25 we analyzed the stabilizationof BSA by some other buffers, such as tris(hydroxymethyl)amino-methane (TRIS), N-[tris(hydroxymethyl)-methyl]-2-aminoethane-sulfonic acid (TES), N-[tris(hydroxymethyl)methyl]-3-aminopropane-sulfonic acid (TAPS), N-[tris(hydroxymethyl)methyl]-3-amino-2-hydroxypropanesulfonic acid (TAPSO), 4-morpholineethane-sulfonic acid (MES), 4-morpholinepropanesulfonic acid (MOPS),and 3-morpholino-2-hydroxypropanesulfonic acid (MOPSO). Thebuffers TRIS, TES, TAPS, and TAPSO are known as the Tris familybuffers, whereas the MES, MOPS, MOPSO, and MOPS–Na areknown as the morpholine family buffers. Thus, it is worthwhileto compare the stabilization of BSA in the presence of thepiperazine family buffers (HEPES, EPPS, and HEPES–Na) withthose of the Tris and the morpholine family buffers. The denatura-tion of BSA in 1.0 M MES, MOPS, MOPSO, and MOPS–Na occurredat 59 1C, 62 1C, 61 1C, and 65 1C, respectively. The denaturation ofBSA in 1.0 M of TRIS, TES, TAPS and 0.7 M TAPSO was observed at62 1C, 60 1C, 58 1C, and 60 1C, respectively. In general, all the three

families of buffers show a strong ability for the stabilization of thenative structure of BSA against thermal denaturation, but theability of the morpholine family buffers is slightly better thanthose of the Tris and piperazine family buffers.

Any change in the native structure of proteins directly relatesto their peptide backbone structure. Therefore, to more clearlyunderstand the influence of the investigated buffers on thepeptide backbone unit and also to consider the role of hydra-tion layer, we experimentally determined the phase transitiontemperature of a thermoresponsive polymer, poly(N-isopropyl-acrylamide) (PNIPAM), in an aqueous solution of the buffers(HEPES, EPPS, HEPES–Na and MOPS–Na) at various concentra-tions (0.0, 0.1, 0.3, 0.5, 0.7, and 0.9 M). In the present study, thehydrodynamic diameters of PNIPAM in 0.0 M to 0.9 M ofinvestigated buffer solutions as a function of temperature weremeasured by DLS. The results of the measurements are graphicallyshown in Fig. 2 and are compiled in Tables S.5 to S.8 in the ESI.†

It can be seen from the tabulated values of dH in Tables S.5–S.8(ESI†) and in the phase transition curves of the PNIPAM in 0.0 Mto 0.9 M HEPES, EPPS, HEPES–Na, and MOPS–Na buffer solutions

Fig. 1 DLS graphs for the hydrodynamic diameter (dH) of BSA in 0.05 M, 0.5 M, and 1.0 M HEPES (a), EPPS (b), HEPES–Na (c), and MOPS–Na (d) buffer asa function of temperature:(-’-) 0.05 M, (-K-) 0.5 M, and (-m-) 1.0 M.

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(Fig. 2) that the LCST value for PNIPAM in pure water is 34 1C,and it substantially decreases with the increasing concentrationof buffers. The phase transition temperatures or the LCST ofPNIPAM in 0.1 M, 0.3 M, 0.5 M, and 0.7 M solutions of HEPES,EPPS, and MOPS–Na buffers are 33 1C, 30 1C, 27 1C, and 24 1C,respectively. At a higher concentration (0.9 M), the LCST ofPNIPAM was observed to be 20 1C in HEPES buffer solution andto be 21 1C in EPPS or MOPS–Na buffer solution. The LCST ofPNIPAM were found to be 32 1C, 28 1C, 24 1C, 19 1C, and 14 1Cin 0.1 M, 0.3 M, 0.5 M, 0.7 M, and 0.9 M HEPES–Na buffersolutions, respectively. Thus, the effect of HEPES, EPPS andMOPS–Na buffers on lowering the LCST of PNIPAM is almostsimilar. However, HEPES–Na buffer remarkably suppressed theLCST of PNIPAM to a lower value, especially at higher concen-trations. The overall tendency of these investigated buffers fordecreasing the LCST of PNIPAM follows the order of HEPES–Na 4HEPES 4 EPPS E MOPS–Na. The enhanced tendency ofHEPES–Na over the other investigated buffers may be attribu-ted to its special structural features and polar nature. On the

structural basis, the only difference between HEPES andHEPES–Na buffers is that the sulfonic hydrogen in HEPES isreplaced by a sodium ion in HEPES–Na. Thus, in an aqueoussolution, the dehydrogenated sulfonic group in HEPES–Naexhibits a strong affinity to water, leading to the phase transi-tion of PNIPAM at a comparatively lower temperature.

A similar behavior for the LCST of PNIPAM was alsoobserved with the morpholine family buffers, i.e., MES, MOPSand MOPSO.23 The LCST of PNIPAM in 0.9 M MES, MOPS, andMOPSO buffer solution were observed to be 25 1C, 24 1C,and 21 1C, respectively. The LCST quenching tendency of thesemorpholine family buffers for PNIPAM follows the orderof MOPSO 4 MOPS E MES. The influence of the piperazinefamily buffers on the LCST of PNIPAM was found to be greaterin comparison with the morpholine family buffers. Thisenhanced tendency is expected due to the presence of an extrahydrogen bond donating hydroxyl group in the piperazinefamily buffers. In addition, the MOPS–Na buffer suppressesthe LCST of PNIPAM to a greater extent compared to the

Fig. 2 Hydrodynamic diameter (dH) for PNIPAM in 0 M, 0.1 M, 0.3 M, 0.5 M, 0.7 M, and 0.9 M HEPES (a), EPPES (b), HEPES–Na (c), and MOPS–Na(d) buffers as a function of temperature: 0.0 M (J), 0.1 M (’), 0.3 M (K), 0.5 M (m), 0.7 M ( ), 0.9 M (.).

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MOPS buffer. This behavior is similar to the trend observed inthe case of HEPES–Na and HEPES buffers, and can be attributedto the same structural basis.

Fig. 3 shows the intensity distribution curves for PNIPAM in0.0 M to 0.9 M HEPES–Na buffer solutions at three differenttemperatures: before the phase transition, at the phase transi-tion, and after the phase transition. To keep the discussionconcise and clear, we do not present the intensity distributioncurves for PNIPAM in the presence of other investigated buffers

(EPPS, HEPES, and MOPS–Na). However, similar trends wereobserved with each buffer. As shown in Fig. 3(a), PNIPAM inpure water has only one peak at 10 1C, corresponding to thehydrodynamic diameter of 18.1 nm. This peak indicates that allthe PNIPAM molecules are in hydrated form at lower tempera-tures. The peak at the phase transition temperature (34 1C)corresponds to the hydrodynamic diameter of 274.7 nm, whichbecomes broader and has less intensity compared to the peak at10 1C. This illustrates that at the phase transition temperature,

Fig. 3 The intensity distribution curves of PNIPAM in various concentrations of HEPES–Na buffer: 0.0 M (a), 0.1 M (b), 0.3 M (c), 0.5 M (d), 0.7 M (e),and 0.9 M (f).

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the amide groups on one chain become available to interactwith the amide groups on a nearby chain, leading to anaggregation in the polymer chains. PNIPAM peaks in 0.1 Mand 0.3 M HEPES–Na buffer solutions (Fig. 3(b) and (c)) followthe similar trends as those appearing in pure water, except forthe size of the particles, which are larger compared to those inpure water, even at lower temperatures. This behavior is expecteddue to the influence of the buffer on PNIPAM molecules. It can beseen from Fig. 3(d–f) that at higher buffer concentrations (0.5 M,0.7 M, and 0.9 M) and at 10 1C, two peaks were observed: the firstpeak corresponds to the hydrated PNIPAM molecules and thesecond peak corresponds to the aggregated PNIPAM molecules.The appearance of the second peak in the aggregated regionindicates that the PNIPAM molecular structure begins to collapseand aggregate in the presence of higher concentrations ofinvestigated buffers, even at lower temperatures. At the respectiveLCST temperature, a single peak of PNIPAM in 0.5 M and 0.7 MHEPES–Na buffer solutions was observed, corresponding to theaggregated globule conformation. However, two lower intensitypeaks were observed in 0.9 M buffer solution. The first peakindicates that some PNIPAM molecules exist in the hydratedform, and the second peak indicates the population of PNIPAMmolecules in an aggregated conformation. At temperaturesbeyond the phase transition, a single peak was observed, repre-senting the aggregated PNIPAM molecules with a dominatingintra-molecular hydrogen bonding network.

Again considering the phase transition curves of PNIPAM,below the phase transition temperature or the LCST point, thesize of the PNIPAM polymer remains unchanged, and thenabruptly increases to a higher value at the LCST point (Fig. 2).This behavior confirmed that the polymer exists in a hydratedrandom coil conformation at temperatures lower than the LCSTvalue and changes into a globular form at LCST. In an aqueousPNIPAM system, two types of intermolecular interactions existsimultaneously. The first is the hydrophobic interaction betweenthe pendent isopropyl groups of the side chain and backbone, andthe second is the hydrogen bonding between the amide groupmoiety and the surrounding water molecules. From the previousinvestigations,106–110 it was disclosed that the transition from acoiled to a globular state is associated with these two types ofinteractions. With increasing temperature, the network of hydro-gen bonding becomes weaker and the amide moiety of onepolymer chain becomes available to interact with the pendentisopropyl group on another polymer chain of PNIPAM, leading toa hydrophobic collapse of PNIPAM. Therefore, the aggregationbehavior of PNIPAM is considerably similar to that of proteins.However, phase transition in the case of PNIPAM is reversible andcan be easily recovered via a cooling process.

Zhang et al.111 studied the effects of the Hofmeister anionson the LCST of PNIPAM as a function of varying anions. Theysuggested that the presence of Hofmeister anions can influencethe LCST of PNIPAM in three ways. First, the anion can polarizethe surrounding water molecules that are involved in hydrogenbonding with the amide group. Second, these anionic speciescan interfere with the hydrophobic hydration of the macromo-lecules by forming hydrogen bonds with water molecules or by

increasing the surface tension of the cavity adjacent to thebackbone and the isopropyl side chain. Third, the anion maydirectly bind to the amide group of PNIPAM. Furthermore, theysuggested that the first and second effects should result in thesalting-out of the polymer, thereby lowering the LCST, and thatthe third effect should lead to the salting-in of the polymer. Inthe present case, the LCST of PNIPAM decreases and causesbuffering-out of the PNIPAM from the investigated buffersolutions, in the similar manner as observed in the case ofinorganic salts. Thus, this indicates that the studied zwitter-ionic buffers are behaving as kosmotropic ions and are stronglyinteracting with the water molecules instead of directly inter-acting with the peptide backbone. The investigated buffer(HEPES, EPPS and HEPES–Na) molecules are polar compoundsand are highly soluble in water. Recently, Taha et al.112 studiedthe mechanism of the phase separation of aqueous organicsolvents in the presence of HEPES buffer via molecular dynamicsimulation. It was found that by the introduction of HEPES bufferto aqueous solutions of tetrahydrofuran (THF), 1,3-dioxolane,1,4-dioxane, 1-propanol, 2-propanol, tert-butanol, acetonitrile, oracetone, the organic solvent can be excluded from water to form anew liquid phase. In the present case, it is most likely that theinteractions of the buffer molecules with the hydrated layer,which surrounds the PNIPAM, are stronger than those withPNIPAM, thus leading to hydrophobic collapse of PNIPAM. Byincreasing buffer concentration, the buffer–water intermolecularinteraction also becomes stronger, thus resulting in the phasetransition occurring at relatively lower temperatures. Thisexplains why the LCST of PNIPAM decreases with the increasein respective buffer concentration. From these results, it might beexpected that these buffer molecules, particularly at high concen-tration, would interact with the hydration layer surrounding theprotein. This causes the proteins to attain a subsidiary globuleconformation. Moreover, several layers having buffer and watermolecules in combination arrange around the hydrationlayer adjacent to the protein. These external combined layers,including the buffer and water molecules, protect the proteinfrom external impetus; in addition, the internal hydrationlayer maintains the structure of the protein in a globularconformation.

FTIR measurements for PNIPAM

FTIR spectra analysis is recognized as a highly sensitive techni-que, not only for the conformational changes in the moleculebut also for the change in the local micro-environment aroundthe molecule, and for the change in the interaction between themolecules, and particularly for the change in the vibrationsof an amide group. Therefore, to confirm the suggested mecha-nism of interactions between PNIPAM and the investigatedbuffers (HEPES, EPPS, HEPES–Na and MOPS–Na) molecules,we made a series of FTIR measurements for the samples ofPNIPAM in pure water and in 0.1 M, 0.5 M, and 0.7 M investi-gated buffer solutions at 20 1C. The results of the measurementsare presented in Fig. 4(a–d). The PNIPAM in aqueous solutionexhibits two peaks, characterized as amide I at 1624 cm�1 andamide II at 1562 cm�1.113 The amide I peak corresponds to the

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CQO stretching mode of vibration and the amide II peak to thecombination of the C–N stretching (40%) and N–H bending(60%) modes of vibration. Thus, these two peaks provideimportant information about the hydrogen bonding of theamide group. It can be seen from Fig. 4(a–d) that no significantspectral changes in the amide I and amide II bands of PNIPAMwere observed in the presence of the investigated buffers.In addition, no extra spectral band appeared due to the intro-duction of the investigated buffers. Thus, the outcomes fromthe spectral measurements reject the possibility of directinteraction between the investigated buffers and PNIPAMmolecules. The band centered at 1624 cm�1 is attributed tothe hydrogen bonding between the carbonyl group and watermolecule ({CQO� � �H–O–H), and the shoulder at 1650 cm�1 isrelated to the hydrogen bonding between the carbonyl groupand amide group, either on the same chain or on the differentchains of the polymer ({CQO� � �H–Nz).114–117 This means thatthe amide I peak provides information about both dehydration, aswell as inter- or intra-cross-linking of the polymer. The amide Ipeak remains unaffected with increase in buffer concentration,

indicating that below the LCST point, the majority of the carbonylgroups remain hydrogen bonded with water molecules in theexperimental concentration range of the buffers. However, theincrease of shoulder intensity with the increase of bufferconcentration may be attributed to the formation of intra- orinter-hydrogen bonds between the carbonyl and amide groups({CQO� � �H–Nz). The gradual shift in the amide II peak froma higher frequency to a lower frequency with increasing bufferconcentration also indicates the conversion of ({N–H� � �H–O–H)bonds into ({N–H� � �OQCz) hydrogen bonds. Therefore, thepresence of buffer causes the dehydration of the amide group,especially at higher buffer concentrations. Consequently, PNIPAMmolecule competes with the buffer molecules for water. When theconcentration of buffers increases at a particular temperature, thewater molecules surrounding the PNIPAM will be withdrawn awayby the buffer molecules. This causes the transition of polymermolecule from a coiled to a globular aggregated conformation,even at low temperatures. Therefore, the outcomes of the FTIRspectra are in accordance with the DLS measurements for thePNIPAM system.

Fig. 4 FTIR spectra of PNIPAM in H2O buffer solution at 25 1C: HEPES (a), EPPS (b), HEPES–Na (c), and MOPS–Na (d): solid black line (0.0 M), solid redline (0.1 M), solid cyan line (0.5 M), and solid blue line (0.7 M).

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The Jones–Dole parameters (A and B)

From the results of the DLS and FTIR measurements forPNIPAM in aqueous buffer solutions, it appeared that theinvestigated buffer molecules interact strongly with the watermolecules. To further confirm the strong interactions betweenthe investigated buffer and water molecules, and to analyze theinherent kosmotropic and chaotropic behavior of these buffers,we obtained A and B coefficients by fitting the viscosity data tothe Jones–Dole equation for each investigated buffer–watersystem. The calculated values of A and B are listed in Table 1.The values of the densities and viscosities used in the calcula-tion of the parameters A and B for each buffer–water system arereported in the Table S.9 (ESI†).

It has been suggested that the value of A represents thebuffer–buffer interaction and the value of B represents thebuffer–water interactions. It has also been reported that the natureand the positive or negative sign of the constant B directlydetermines the kosmotropic (structure maker) or the chaotropic(structure breaker) nature of the solute. The positive value ofB represents the kosmotropic nature of the solute, whereas the

negative value of B represents the chaotropic nature of thesolute.70 From the listed values of the constants A and B foreach investigated buffer in Table 1, it can be seen that the valueof B is positive and very high compared to the value of A. Thisclearly indicates that the dominating interaction in the solu-tion is the buffer–water interaction and all these buffers exhibitkosmotropic nature. Previous researchers118–120 have mentionedthat kosmotropes generally interact strongly with water molecule incomparison to the interaction of water molecules among them-selves. Therefore, kosmotropes usually influence the hydrationlayer in the close proximity of the protein, reducing the entropyof their vicinal water, and thus usually act as protein structurestabilizers. Moelbert et al.121 reported a similar protective effect

Fig. 5 The UV-visible absorption spectra of BSA in various concentrations of buffers at pH = 7.0 and T = 25 1C: HEPES (a), EPPS (b), HEPES–Na (c), andMOPS–Na (d); (—), 0.05 M; (––), 0.5 M; (� � �), 1.0 M.

Table 1 Jones–Dole equation parameters for the biological buffersHEPES, EPPS, HEPES–Na, and MOPS–Na in aqueous solutions at 25 1C

Buffer A B

HEPES �0.146 0.830EPPS �0.154 0.968HEPES–Na �0.305 1.435MOPS–Na �0.132 0.863

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of the kosmotropic sugar molecules for the protein againstdenaturation induced by external factors, such as heating orcooling. In addition, it has also been suggested that the largekosmotropic molecules, due to their differential interactionbehavior, are excluded from the vicinity of a protein, leadingto the preferential hydration of the protein.122,123 Thus, ourexpectation about the mechanism of the stabilization of protein inaqueous solutions of these kosmotropic buffers is in accordancewith the previous observations.

UV-visible measurements

To examine the interactions of the protein BSA with the investi-gated buffers (HEPES, EPPS, HEPES–Na and MOPS–Na), UV-visibleabsorption spectra were measured for BSA in 0.05 M, 0.5 M, and1.0 M of buffer solutions (pH = 7) at 25 1C. The absorption spectrawere measured in the range of 200–350 nm, and the results arepresented in Fig. 5(a–d). It can be seen from these graphs thatthe absorption spectra of the protein BSA in aqueous solutionsof buffers have two distinct peaks. The aqueous solutions of theprotein BSA on irradiation with UV light exhibit two clear peaks,the first in the short wavelength region (200–250 nm) andthe second in the higher wavelength region (255–320 nm).124

Generally, the structure of BSA contains polypeptide chainsand aromatic amino acid residues, such as 20 tyrosine (Tyr),27 phenylalanine (Phe), and two homologous tryptophan residues(Trp 212 and Trp 134). Kuipers and Gruppen125 suggested that themolar excitation coefficient of tryptophan residue is 30 timeshigher than that of the peptides, whereas those of phenylalanineand tyrosine residues are six times higher than that of the peptidegroup of proteins. Thus, the first peak of BSA in the lowerwavelength region is considered to be the characteristic of thepolypeptide backbone structure CQO, and results from thep- p* transition.126 The second absorption peak in the higherwavelength region is considered to be characteristic of thearomatic amino acids residues (Trp, Tyr, and Phe), and resultsfrom the n - p* transition.126

As shown in Fig. 5(a–d), the maxima of the first peak, in thelower wavelength region of BSA in 0.05 M HEPES and MOPS–Nasolutions is exhibited at 238 nm, whereas those of BSA in 0.05 MEPPS and HEPES–Na buffer solutions appear at 240 nm. Thispeak shifts to 241 nm for the BSA in 0.5 M MOPS–Na buffersolution and to 242 nm for the BSA in other investigated buffersolutions (HEPES, EPPS and HEPES–Na). In the higher concen-trations of the buffer solutions (1.0 M), the maxima of the firstpeak shifts to 242 nm, 243 nm, and 244 nm for MOPS–Na,HEPES or HEPES–Na, and EPPS systems, respectively. Overall,with the increase of buffer concentration, the peak of BSAcorresponding to the amide group shifts to a higher wavelengthand becomes less intense. Polet and Steinhardt127 mentionedthat the amide group of proteins, which is exposed to theaqueous surroundings, undergoes p–p* transition on ultra-violet irradiation. Recently, Sandhya et al.128 suggested that ared shift of the peak of BSA in the lower wavelength region mayoccur due to the change of the micro-environment around thepeptide moieties by highly polar solvents. Thus, the observedbathochromic shift with increasing buffer concentration,

as seen from Fig. 5(a–d), might be the result of the disturbancein the surroundings of the polypeptide caused by the inter-action of the investigated buffers with water molecules. Thisassumption is supported by the observed mechanism of thecollapsing of polymer PNIPAM in the presence of the investi-gated buffers, even at lower temperatures. We also suggest thatthese buffers strongly interact with water and destroy thehydration structure around the polymer, and thus result in theaggregation. Certainly, these highly polar buffers (HEPES, EPPS,HEPES–Na, and MOPS–Na) behave in a similar manner with BSAand strongly interact with the water shell over the peptide groupof BSA. This provides shielding to the peptide group fromaqueous environments and lowers the energy of the p and p*electron clouds. Thus, this facilitates the low-energy p–p* transi-tion and results in the observed bathochromic shift.

All the maxima of the second peak of BSA in HEPES, EPPS,HEPES–Na, and MOPS–Na buffer solutions appear at 280 nm(Fig. 5(a–d)). No shifting was observed for this peak of BSA in allthe investigated buffer solution over the entire concentrationrange (0.05 M, 0.5 M, and 1.0 M). In addition, with increase inbuffer concentration, minor variations in the intensity of thispeak were noticed. The minor changes in the intensity of thispeak may be due to the variation in the micro-environmentaround the aromatic amino acid residues.

Fluorescence measurements

Fluorescence technique has been proven to be very useful inthe investigation of molecular environment in the vicinity ofchromophore molecules. Because, the fluorescence intensity ofa fluorophore molecule is highly sensitive to the proximityof the other surrounding molecules.10,11 Therefore, to confirmthat the investigated buffers produce disturbance in the sur-roundings of the peptide moiety of BSA, and to further checkthe overall effect on the structure of BSA due to the presence ofthe respective buffer molecules, we measured the emissionspectra of BSA in 0.05 M, 0.5 M, and 1.0 M HEPES, EPPS,HEPES–Na, and MOPS–Na buffer solutions (pH = 7) at excita-tion wavelengths of 230 nm and 280 nm. The results of themeasurements are presented in Fig. 6(a–d) at 230 nm and inFig. 7(a–d) at 280 nm. Sandhya et al.128 experimentally deter-mined the 3D fluorescence spectra of BSA. They reported that thefluorescence peak at the excitation/emission of 230 nm/340 nmin the 3D spectra appeared mainly due to the peptide backbonestructure of the protein, and the fluorescence peak at theexcitation/emission wavelength of 280 nm/340 nm was char-acteristic of aromatic amino acid residues (Phe, Tyr, and Trp).It can be clearly observed from Fig. 6(a–d) that upon excitationat 230 nm, the fluorescence intensity of BSA is significantlyquenched in the presence of investigated buffer molecules. Theobserved quenching effect increases with increasing the respec-tive buffer concentration. It can be directly expressed from theresults that the investigated buffers (HEPES, EPPS, HEPES–Na,and MOPS–Na) strongly influence the peptide moiety of BSA.The fluorescence peaks of BSA, upon excitation at 280 nm, showthat no significant changes were found in HEPES and EPPS buffersolutions, as shown in Fig. 7(a) and (b), respectively, over the

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entire concentration range. However, a marked quenching behaviorwas observed from MOPS–Na- and HEPES–Na-containing systems,Fig. 7(c) and (d), respectively, especially at higher concentrations.Probably, these buffers exclude the water molecule from the vicinityof protein. The exclusion of the water molecule from the vicinity ofprotein may change the micro-environment around the amidegroup, as well as around the aromatic amino acid residues, andlead to the observed changes in the intensity of fluorescence peak.Thus, the results of the fluorescence measurements strongly sup-port the finding from the UV-vis spectra. HEPES–Na and MOPS–Nabuffers influence the surroundings of BSA to a slightly greater extentdue to their special structural features and polar nature. This mightbe the reason that these buffers appear to be better stabilizers forBSA over HEPES or EPPS. Moreover, no shifting was found inthe fluorescence spectra of BSA in the presence of any investi-gated buffers, neither at 230 nm nor at 280 nm, indicating thatthe structures of BSA remain in the native state due to theinfluence of these buffers.

FTIR measurements for BSA

To further confirm the influence of these buffers on the amidemoiety of the protein BSA and the adverse effect of these inter-actions on the structure of the protein, we obtained the FTIR

spectra of BSA in 0.05 M, 0.5 M, and 1.0 M HEPES, EPPS, HEPES–Na,and MOPS–Na buffer solutions at pH = 7. The spectrum wasrecorded at 25 1C and in the range of 1700–1350 cm�1. The resultsof the measurement are presented in Fig. 8(a–d). For comparisonpurposes, the FTIR spectra of BSA in 1.0 M HEPES, EPPS,HEPES–Na, and MOPS–Na buffer solutions (pH = 7) are pre-sented in Fig. 8(e). The IR spectrum of the protein shows twodistinct bands: amide I, (mainly due to the CQO stretchingmode of vibration) and amide II (due to the coupling of theN–H bending and C–N stretching modes).129 As can be seen fromFig. 8(a–d), no significant changes occur in these amide intensitybands of BSA due to the presence of any of these examined buffers,over the entire concentration range (0.05 M, 0.5 M, and 1.0 M).This indicates that the possibility of direct interactions betweenthe buffers and BSA molecule is negligible. However, the observedchanges in the UV-visible and fluorescence spectra of BSA in theinvestigated buffer solutions, especially at higher concentrations,might result from the change in the conformation of the proteincaused by the influence of these buffers on the surroundinghydration layers of the protein. In addition, neither bathochromicnor hypsochromic shifts were observed in the IR spectra of BSA inthe presence of the investigated buffers, indicating that the buffermolecules support the structure of the protein.

Fig. 6 Fluorescence spectra of BSA (at lex = 230 nm) in 0.05 M, 0.5 M, and 1.0 M buffers (pH = 7.0): HEPES (a), EPPS (b), HEPES–Na (c), and MOPS–Na(d) at 25 1C; (-’-) 0.05 M, (-K-) 0.5 M, and (-m-) 1.0 M.

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Molecular docking studies

To explore the possible binding sites and the preferred orienta-tion of BSA with HEPES, EPPS, HEPES–Na, and MOPS–Nabiological buffers, we performed the molecular docking calcu-lation for the BSA in the presence of the investigated buffers bythe Molegro virtual docking program.88,89 The most likely bind-ing sites and the optimal conformation of BSA were determinedon the basis of the highest ranking docking score. The results ofthe docking calculations are presented in Fig. 9(a–d). It can beclearly seen from these graphs that the investigated buffers havehydrogen-bonding interactions with the amino acid residues indomains II and III of BSA. Previous research46 also found thatthe potential binding sites for the polar ligands in protein BSAexist in domains II and III of the BSA. On the basis of thedocking calculations, the residues found adjacent to the HEPESbuffers are Lle 387, Asn 390, Leu 406, Arg 409, Leu 429, Val 432,Cys 436, Arg 444, Cys 447, Thr 448, and Leu 452 (Fig. 9(a)). TheHEPES buffer was found to form 8 hydrogen bonds with theseamino acid residues. As shown in Fig. 9(b), the residues foundadjacent to the EPPS buffer are Asn 390, Cys 391, Leu 406,

Arg 409, Tyr 410, Leu 429, Val 432, Gly 433, Arg 435, Cys 437, Arg444, Cys 447, Thr 448, and Leu 452. The EPPS buffer forms ninehydrogen bonds with these amino acid residues. Fig. 9(c) showsthat the residues Asn 390, Cys 391, Gln 393, Phe 394, Phe 402,Leu 406, Arg 409, Tyr 410, Leu 429, and Leu 452 exist nearHEPES–Na. The HEPES–Na buffer was found to form eighthydrogen bonds with these amino acid residues. The residuesfound adjacent to the MOPS–Na are Lys 204, Cys 460, Leu 462,His 463, Glu 464, Lys 465, Cys 476, and Thr 477 (Fig. 9(d)). TheMOPS–Na was found to have only five hydrogen bonds withthese amino acid residues. The fewer hydrogen bonds in thecase of MOPS–Na compared to the piperazine family bufferswas expected due to the absence of an extra hydroxyl group inMOPS–Na. One thing we want to highlight is that the dockingcalculations were performed for the mixtures in the gaseousphase. Thus, in the present aqueous case, the existence of suchobserved hydrogen bond interactions are always doubtful. How-ever, even if these direct interactions between the buffer and BSA inaqueous solution occur, they certainly stabilize the structure of theprotein, as indicated by the results of the DLS, FTIR, and fluores-cence measurements. From the docking calculations, our main aim

Fig. 7 Fluorescence spectra of BSA (at lex = 280 nm) in 0.05 M, 0.5 M, and 1.0 M buffers (pH = 7.0): HEPES (a), EPPS (b), HEPES–Na (c), and MOPS–Na(d) at 25 1C; (-’-) 0.05 M, (-K-) 0.5 M, and (-m-) 1.0 M.

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was to locate the potential binding sites in the protein BSA with theinvestigated buffers. Proteins are expected to coexist with buffers.Therefore, these findings of the docking study can be very helpfulfor drug development and for pharmaceutical industries.

Activities of a-CT

The biological activities of biomolecules, such as enzymesor proteins, are directly related to their active conformation.

The basic principle by which an enzyme catalyzes the reaction isbased on the lock and key hypothesis. The enzyme is only activeor is only able to catalyze a particular reaction only when it is inits active form because any change in its native conformationcan restrict the enzyme from binding to the substrate, and thusrestrict the catalysis of the respective reaction. Thus, by monitor-ing a suitable chemical reaction catalyzed by an enzyme in thepresence of a cosolvent, we can directly evaluate the influence of

Fig. 8 FTIR spectra of BSA in H2O buffer solutions at 25 1C: HEPES (a), EPPS (b), HEPES–Na (c), and MOPS–Na (d): solid black line (0.05 M), solid red line(0.5 M), solid blue line (1.0 M). (e) FTIR spectra of BSA in 1.0 M buffer (pH = 7.0), solid black line (HEPES), solid red line (EPPS), solid blue line (HEPES–Na),and solid green line (MOPS–Na) at 25 1C.

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Fig. 9 Molecular docking of BSA with HEPES, EPPS, HEPES–Na, and MOPS–Na buffers: HEPES (a), EPPS (b), HEPES–Na (c), and MOPS–Na (d).

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the cosolvent on the structure of enzyme. Therefore, to furtherconfirm the fact that the studied buffers (HEPES, EPPS, HEPES–Na,and MOPS–Na) support the native structure of the biomoleculesand provide compact stable conformations, we analyzed the effectof these buffers on the catalytic activity of enzyme a-chymotrypsin(a-CT) for the aqueous hydrolysis of p-nitrophenylacetate (PNPA).The hydrolysis of p-nitrophenylacetate (PNPA) catalyzed by a-CT iswidely recognized and has attracted a significant attention fromenzymologists worldwide.130–132 In addition, the enzymatic rateenhancement of reactions has attracted considerable attention ofchemists in recent years.133–136 The selected enzyme a-CT is a chiefserine protease and is widely distributed in the nature. Enzymea-CT is widely used in industries for catalytic applications.93

Therefore, the enhancement of the catalytic activity of enzymea-CT is highly desirable to achieve a faster and more economicalprocess.

To examine the expected effect, the activities of enzyme a-CTin 0.05 M, 0.5 M, and 1.0 M aqueous solutions of buffers,HEPES, EPPS, HEPES–Na, and MOPS–Na at 400 nm, at pH = 8.0and at T = 25 1C were measured for a time period of 3 minutes.The results of the measurements are presented in Table 2. Fromthe tabulated values of the activities in Table 2, it can be seenthat the biological buffer enhances the catalytic activity of enzymea-CT and makes the enzyme super-active, especially at higherconcentrations. The effect of the super-activity is found to increasewith increasing concentrations of the respective buffer. Theobserved results indicate that the investigated buffers are notonly suitable for maintaining a suitable pH range for the efficientworking of biomolecules, but are also capable of providing a moreactive conformation, and thus it could support the native struc-ture of proteins. Our expectations are in accordance with theresults of Castro,137 who concluded that the optimum activityof a-CT in aqueous solution at pH = 8 is caused due to thecompact native conformations.

Conclusions

The main objective of the present work was to find a systemwith a high possibility of protecting the native structure of theprotein BSA in aqueous solutions against thermal denaturationby means of commonly used biological buffers. Therefore, thethermal denaturation processes of BSA in 0.05 M, 0.5 M, and1.0 M HEPES, EPPS, HEPES–Na, and MOPS–Na biological buffersolutions were analyzed by measuring hydrodynamic diameter(dH) at various temperatures by dynamic light scattering (DLS).

These buffers were found to support the native structure of BSAagainst thermal denaturation. In particular, the denaturation ofBSA occurred at 65 1C in 1.0 M of MOPS–Na solution. This value isconsiderably higher than that of BSA in water (55 1C). To confirmthe role of hydration layer in the stabilization of the protein, weanalyzed the phase transition of the thermoresponsive polymerPNIPAM in 0–0.9 M of the investigated buffer solutions. Theseresults reveal that the hydration layers play a very importantrole in the protection of the protein structure against thermaldenaturation. To explore the mechanism of the interactions ofthese biological buffers (HEPES, EPPS, HEPES–Na, and MOPS–Na)with BSA, we measured the UV-visible absorption, fluorescence,and FTIR spectra of BSA in various buffer solutions at differentconcentrations. It was found that these buffers mainly interactwith the hydration layers surrounding the peptide backbone ofBSA protein. To locate the possible interaction sites between thesebuffers and BSA protein, we conducted molecular docking calcu-lations. The results from the docking studies show that the mostpreferable interacting sites of the protein with the buffers in thegaseous phase are the domains II and III of BSA. In addition, thesuper-activity of the commercial enzyme a-CT in aqueous solu-tions of the investigated buffers was also found. Because thebuffer is an essential component in protein mixtures, the stabili-zation of proteins by these buffer molecules can avoid using anyextra additive in the systems. As a consequence, this series ofstudies is helpful in the understanding and development of moreuseful buffers in biological field.

Funding sources

The authors gratefully acknowledge the financial support fromNational Science Council, Taiwan, through NSC102-2811-E011-014and NSC 102-2221-E011-137-MY3.

Acknowledgements

The authors thank Dr Ho-mu Lin for valuable discussions.

Notes and references

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Table 2 Catalytic activity of a-CT in 0.05 M, 0.5 M, 1.0 M HEPES, EPPS,HEPES–Na and MOPS–Na buffers for the time period of 3 minutes, at400 nm, pH = 8.0, and T = 25 1C

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Buffers more than buffering agent: introducing a new class of stabilizers for the protein BSA

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