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Chemical Physics 307 (2004) 285–290

Towards measurement of the time-resolved heat release ofprotein conformation dynamics

Jason Puchalla a, Daniel Adamek b, Robert Austin c,*

a Department of Molecular Biology, Princeton University, Lewis Thomas Laboratories, Princeton, NJ 08544, USAb AZ Technology at NASA Laboratory for Structural Biology, Code SD46, NASA, MSFC Huntsville, AL 35812, USA

c Department of Physics, Princeton University, 122 Jadwin Hall, Princeton, NJ 08544, USA

Received 4 May 2004; accepted 2 September 2004Available online 28 October 2004

Abstract

We present a way to observe time-resolved heat release using a laminar flow diffusional mixer coupled with a highly sensitiveinfrared camera which measures the temperature change of the solvent. There are significant benefits to the use of laminar flow mix-ers for time-resolved calorimetry: (1) The thermal signal can be made position and time-stationary to allow for signal integration; (2)Extremely small volumes (nl/s) of sample are required for a measurement; (3) The same mixing environment can be observed spec-troscopically to obtain state occupation information; (4) The mixer allows one to do out of equilibrium dynamic studies. The hope isthat these measurements will allow us probe the non-equilibrium thermodynamics as a protein moves along a free energy trajectoryfrom one state to another.� 2004 Elsevier B.V. All rights reserved.

1. Introduction

A central concept in protein folding is that there ex-ists a free energy landscape, that is, a surface which isa multi-dimensional function of all the parameters thatdescribe the conformation of the protein [1]. The energylandscape picture says that as the protein folds it movesover some free energy landscape biologically inactiveconformations to a global minimum which is a collec-tion of conformations which characterize the biologi-cally active ‘‘state’’ of the protein. If we knew theentire surface of this landscape, we could compute thelocal and global minima of the various conformationsthe protein can have. Unfortunately a protein has fartoo many degrees of freedom for such a landscape tobe calculated. Each amino acid in a polypeptide chain,ignoring side chain conformations, can be simplified to

0301-0104/$ - see front matter � 2004 Elsevier B.V. All rights reserved.

doi:10.1016/j.chemphys.2004.09.006

* Corresponding author. Fax: +1 609 258 1115.E-mail address: austin@princeton.edu (R. Austin).

2 rotational degrees of freedom and 1 additional degreeof freedom to account for relative bond angles. Thereby,a typical 100 amino acid protein could be viewed as hav-ing 300 degrees of freedom, making for an impossiblycomplicated energy landscape. This extreme complexitygave rise in the classic note by Levinthal called ‘‘How tofold gracefully’’ [2], where it was pointed out that onsuch a complex landscape it would take an absurdlylong time for a protein to find the lowest valley in thisincredibly complex space if the search was random.Thus, naively proteins should not be able to fold intothe active set of conformational states since there arefar too many conformational states to search through.

Of course, the view that proteins cannot fold assumesthat the free energy landscape of the protein conforma-tional space is flat and that the search for the globalminimum is a diffusional random walk with little bias.Wolynes et al. [3,4] in seminal papers have discussedhow the free energy landscape must be strongly biased.There must exist what can be called funnels which

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strongly bias the path of an unfolded protein towards aglobal free energy minimum in spite of the vast numberof states that a protein can in principle have. A deeperissue is the general validity of a central concept thatcame out of the classic work of Anfinsen [5] called the‘‘Thermodynamic hypothesis’’: the folding process isdriven entirely by the free energy of the conformationof the protein and hence the native, active conformationof the protein is located at the minimum of the Gibbsfree energy of the system. However, anybody who hasever worked in molecular biology is rather suspiciousof this comforting statement: the large complex multi-domain proteins such as RNA polymerase are not stableand, with time, convert to non-functional conforma-tions. Typically the careful investigator stores a complexprotein at �80 �C, but even that condition is not enoughto guarantee full biological activity over the period ofyears. More recently the subject of the epigenetic inher-itance of protein conformations from mother to daugh-ter cells [6] and the realization that even small proteinscan exist in two very different metastable configurations[7] has served to emphasize that there is much more tounderstanding protein conformation dynamics thanstudies of small folding proteins near the thermody-namic minimum in free energy.

While the near-equilibrium (thermodynamic) valuesof the enthalpy, entropy, and Gibbs free energy changesobtained from standard calorimetry [8] yield some infor-mation about the complexity of the free energy surfaceover which a protein moves as it moves between states,only under very limited circumstances has it been possi-ble to directly probe the temporal evolution of the freeenergy of a complex system as it moves between states[9–11]. The flow calorimetry technique presented herecan be used to measure the temporal dynamics of theheat of reaction initiated by a fast mixing process. Atthe same time as the heat evolution is measured, wecan measure the spectroscopic states the system is pass-ing through. If successfully applied, this method wouldprovide a method to examine the dynamics of internalenergy and entropy flow during complex macromolecu-lar reactions and correlate that with state populations.Traditional calorimetric methods are generally restrictedto determining the time-averaged values of various ther-modynamic parameters at or near conditions where theGibbs free energy change between two states is zero.Typically, this involves adding salts or raising the tem-perature to the denaturation point. However, in vivothe dynamics of protein conformational changes in abiological network involve evolution along a complexfree energy surface from a place far from equilibrium.Since the free energy surface of a protein is a complexfunction of many parameters it is entirely possible thatthe folding path traced dynamically far from equilib-rium is quite different from the path followed underdenaturing conditions.

In fact, while many proteins are able to fold sponta-neously and quickly (microsecond time scale) to theirnative set of conformations, many other proteins arenot able to correctly fold spontaneously. In these cases,the enigma of how proteins fold remains a real problem.Nature has evolved a class of proteins called chaperoneswhich guide such proteins to their correct native set ofconformations. As is well known from ideal gas analysisof the Carnot cycle, lowering the entropy of a systemcannot occur spontaneously. Heat DQ must be addedto a system to lower entropy. One might guess that achaperone would need a source of energy in order toguide a protein from many possible conformations toa few, just as a Carnot engine run in reverse (a refriger-ator) needs to have work done on it in order to lower theentropy of the lower temperature heat bath. Of course,since a protein is not an ideal gas, the formulas derivedfor the heat exchange in the Carnot cycle do not apply,but the basic concept of energy input to lower entropy isclear.

Probably the most beautiful and well studied of suchprotein folding machines is the GroEL/ES folding com-plex [12,13]. The Escherichia coli GroEL/ES chaperonincomplex consists of two parts: the GroEL and theGroES subunits. GroEL is a large (14 · 60 kDa) proteinconsisting of 14 identical 60 kDa stacked rings, whileGroES is a smaller (7 · 10 kDa) set of rings which actsas a ‘‘cap’’ to the barrel shaped GroEL complex. Theprotein to be folded resides in the center of theGroEL/ES capped cylinder (Fig. 1). Since loweringthe entropy takes energy, GroEL/ES binds the energyrich ATP molecule to the equatorial domains. The bind-ing of ATP results in the hydrolysis of ATP to ADP, aconformational change of the GroEL/ES complex andthe release of a folded protein from the interior of thecylinder. The binding of ATP and its subsequent hydrol-ysis results in the release of chemical energy which canbe diverted to either heat release if no protein is in thefolding barrel or the folding and release of a protein ifthe GroEL/ES complex is occupied. Surprisingly, ATPhydrolysis seems only required for the release of ligandsand not fold initiation.

2. A time-resolved calorimetric diffusional mixer

In general, we wish to conduct a series of time-resolved measurements including: (1) the heat releasedDQ(t) into the solvent surrounding the protein moleculesduring protein conformational changes to establish theenthalpy changes DH(t); (2) the volume changes DV(t)to establish the work done on the protein and thusestablish the internal energy changes DE(t); and (3) thespectroscopic state changes to obtain the Gibbs energychanges DG(t).

Fig. 1. A side view showing the GroEL barrel and dome-shapedGroES. Each GroEL donut is composed of a ring of seven subunits.The amino acid residues are color-coded in order of the Debye–Wallertemperature factors. Blue corresponds to the smallest temperaturefactor and hence the smallest thermal motions. Taken from [14]. (Forinterpretation of the references to color in this figure legend, the readeris referred to the web version of this article.)

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The most frequent approaches to initiate foldingreactions in kinetic experiments have been rapid mixingtechniques in either stopped-flow or continuous-flowoperation mode [15–18]. If we could watch a proteinevolve from a presumably random unfolded structureto a native-folded state and know exactly the shape,internal energy and entropy at every step along theway, we would go a long way towards understandinghow proteins fold. In order to do this we need to startwith a well-defined population of molecules which arein some basin of states (say, the molten globule state)and change conditions rapidly enough so that it is pos-sible to watch them move along the free energy surfaceto the new basin. The most general way to do this isto suddenly change the environment of the solvent sur-rounding the protein so that the free energy surfaceshifts to a topology, such that a new state now lies lowerthan the basin that the protein is presently occupying.The protein will now start to move along the path ofsteepest descent (the gradient of the free energy surface)towards the new local basin of minimum free energy.

All of this requires rapid mixing of fluids. Mixing isdriven by diffusion, and is therefore slow over macro-scopic length scales since x2 = 2Dt. It is frustrating,for example, to wait passively for cream to mix evenlyinto a cup of coffee without stirring. Many liquid-phase

chemical and biological processes exhibit dynamics thatcannot be resolved in reaction kinetics experiments be-cause they are faster than the mix times of conventionalmixers. Protein folding is one important example inwhich the current emphasis is on time scales shorter thanthe millisecond mixing times attainable with traditional,stopped-flow methods. The time for diffusion to mix avolume can be reduced by the creation of chaotic turbu-lence, as when coffee and cream are stirred with a spoon.Similarly, the fastest mixers used for reaction kineticsintroduce turbulence by forcing reactant streams at highvelocity through a nozzle. This technique can yield mixtimes below 100 ls but it consumes large volumes (10�sof ml) of sample. The transit time of the mixed fluidthrough the nozzle further imposes a dead time duringwhich the reaction is obstructed from view. A fasteralternative to turbulent mixing is to reduce the lengthscale over which the fluids must diffusively mix: in theparlance of the coffee metaphor, to use a very smallcup. We have previously described and used a microflu-idic, continuous-flow mixer capable of diffusive mixingtimes of less than 10 ls [15]. This was achieved withoutinducing turbulence or chaos [19,20] and in an openarchitecture that reduces dead time by permitting obser-vation of the entire mixing process.

Our previous diffusional mixer worked only with verysmall volumes which are surrounded by a much largervolume of fluid. As we discussed above, in order toexamine the thermodynamics of the system we need tomeasure the heat flow DQ into the solvent from the pro-tein and the volume change DV. Both of these are extrin-sic quantities. The total heat released into the solventand the total volume change of the protein scales asthe relative fraction of the volume of the sample beingmixed. Hence, we had to design and build a new mixerwhich could mix large volumes quickly. The design wecame up with was to use through-wafer etching to pro-duce multiple streams of fluid side by side, so that eachstream could mix via diffusion with a much smaller vol-ume (Fig. 2). Parasitic heat flow into the surroundingetched structure will be a problem. One way around thisis to isolate the mixing part of the wafer by ‘‘back etch-ing’’ the mixing and flow part of the wafer so as to re-duce the mass of the supporting structure.

We have recently succeeded with the assistance of Pe-ter Galajda in making such a mixer. Fig. 3 shows amicrophotograph of the structure, showing how the wa-fer was through-etched to provide side-by-side mixingcapability of 35 channels.

3. Initial tests of the heat-release diffusional mixer

The time-dependent volume change DV(t) for pro-teins from a denatured configuration to a folded config-

Fig. 2. The basic design of a large volume diffusional mixer. Fluid Aenters from the right along an etched pathway, fluid B enters from belowon the back side of the chip andmixes by diffusion as both fluidsmove tothe right. In the schematic shown here both fluids enter at the samepressure. In the actual chip discussed in Fig. 3 the two fluids are keepseparate until further down to balance hydrodynamic impedances.

Fig. 3. (a) The through-wafer etched structure, showing the region ofthe fluid A flow and the fluid B flow from below. This image is topilluminated. (b) Same view with back illumination, showing how the Bfluid ports are etched entirely through the wafer. The input rectanglesare 10-lm wide and 100-lm long. (c) Mixing region were the A and Bfluids are allowed to contact and mix by diffusion. The walls are 1-lmthick and the fluid regions are 5-lm wide.

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uration, critical for extracting the internal energychanges E from the enthalpy H, has never been meas-ured and so we will pass at this point concerning what

sort of values it could have [21–23]. The enthalpy changeH which gives rise to the heat release is also a trickynumber. In thermodynamics, one is only concerned withthe initial and final free energy levels of the molecule,not the height of the barriers between the states. Thus,the enthalpies and entropies of the kinetic barriers thatconnect states are not the numbers we can use for eval-uation of the probable value of the released heat. Whilethe time-dependent heat release and volume changes de-tail the kinetics, in the end the final temperature differ-ences integrated along the path are independent of thesize of the barriers.

In our case, since we measure the spontaneous heatreleased (or absorbed) by the protein as it folds we meas-ure not the specific heat of the protein Cp by feedingheat into the system and observing the temperaturechange of the system, but the reverse: we measure thechange in temperature DT of the surrounding solventdue to heat released by the protein as it folds. The spe-cific heat of the solvent can be assumed to be a constantfor the very small temperature changes expected sincethe protein represents at most 1% of the volume of thesample.

We can do some rough estimates of the net tempera-ture rise that can be expected in time-resolved thermody-namics after the reaction has gone to completion. Wecan guess that at 20 �C the approximate value for theenthalpy change DH to fold a small protein such as myo-globin or lysozyme of molecular weight about 20,000 g/mole will be about 80 kJ/mole [24]. At a concentrationof 10 mg protein/ml of solvent the molarity of the pro-tein is about 5 · 10�4 moles/l. Since the specific heat ofwater is 4.2 J/ml K, we can estimate that unfolding a 10mM solution of protein will raise the temperature of thesolvent about 100 mK. Since the surface temperaturewill vary with the distance down the channel as the pro-tein folds ultimately increasing by 100 mK, an image ofthe mixer surface in the IR provides a measure of heatrelease versus time. This can be done with the new gen-eration of cryogenically cooled bolometer cameraswhich can stare at the surface of an object in the mid-IR (3–5 lm wavelength) and measure the radiated en-ergy. We have access, through NASA Marshall SpaceFlight Center, to an Indigo Phoenix liquid nitrogencooled 320 · 256 element InSb array camera. The cam-era has spectral bandpass of 3–5 lm, and a NETD of20 mK per frame implying that integrating for 400frames can achieve a resolution of 1 mK per pixel, wellin excess of the expected 100 mK temperature changesexpected for a 10 mM folding/unfolding experiment.

In order to test the performance of this camera wehave carried out a simplified experiment. 1-propanolwas diffusively mixed with water in a simple T-mixeras discussed in [17]. The mixer was machined out of ac-rylic to minimize heat conduction losses. The dilution of1-propanol in water is known to raise the temperature of

Fig. 5. Profiles of the temperature vs. distance in the channel of Fig. 4at three flow speeds.

Fig. 6. Temperature rise observed upon mixing 24 lM GroEL/60 lMGroES with 5 mM ATP. The flow was 3 nl/s, yielding an average flowvelocity of 3.5 mm/s in the 125 micron wide by 75 micron deepchannel. Mixing occurs 0.5 mm from the left side of the device wherefluid is injected.

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the final mixture. In our case, the center channel of themixer contained 50% 1-propanol in distilled water. Theouter two channels had pure distilled water, and the pres-sure on all inputs was the same. Under these conditionsthe known temperature rise of the solution is 500 mK.We used the Indigo camera to observe the chip surface.Fig. 4 shows the image of the mixing solution as a func-tion of the flow speeds. The speeds were varied byincreasing the pressure uniformly in all three channels.Fig. 5 shows a profile of the temperature in the mixingchannel as a function of flow speed. At low flow speedsthe conductivity of the structure leads to a decrease inthe temperature with distance (time) as the liquid movesdown the channel, but as the speed of the flow increasesthe mixing reaction is not completed until farther downthe channel and hence the cooling profile becomes flat-ter. At the fastest speed the fluid velocity is approxi-mately 3 mm/s.

The data shown in Fig. 5 is a single frame from thecamera, yet analysis of the data shows that we are al-ready at a noise level of about 20 mK as per the cameraspecs. Since the flow mixer allows one to sit and stare atthe mixing process for as long as one likes, signal inte-gration may reduce the noise below 1 mK.

As a further test of this technology, and to push thetechnology as far as we can go at present, we have triedto measure a critical aspect of the folding action ofGroEL: the release of heat by the hydrolysis of ATP.We study here an ‘‘empty’’ GroEL/ES complex whichdoes not contain a protein in the central cavity. Wewould expect that if the GroEL/ES complex containsa protein guided along a folding pathway that the freeenergy changes involved in guiding the protein wouldbe reflected in changes in the heat released to the exter-nal solvent.

Thus, we simply mix ATP with a GroEL/ES complexwhich does not contain a protein to be folded. In thiscase we expect that the hydrolysis of the ATP will resultin the release of heat to the solvent. This experiment is a‘‘real life’’ test in that the concentration of the GroEL/ES must be kept below about 50 lM to avoid proteinaggregation (50 lM = 20 mg/ml), so the experimentwas run at 24 lM GroEL, 5 mM DTT, 60 lM GroES,50 mM HEPES buffer, 5 mM KO(Ac), and 10 mMMg(OAc)2 (center channel). The outer two channelscontained 5 mM ATP, 50 mM HEPES buffer, 5 mMKO(Ac), and 10 mM Mg(OAc)2. The reaction we meas-

Fig. 4. Thermal profile of the heat released by mixing 50% 1-propanolin distilled water. These are single frame captures at video rate.

ure here is the binding of ATP to GroEL in the presenceof GroES. We can roughly estimate the expected heatreleased if 20 lM of ATP binds to the GroEL complexand is hydrolyzed to ADP. This reaction is known toyield 7.5 kcal/M (or 31 kJ/M). We would then expectthat the surrounding solvent should warm by 0.15 mKfor the turnover of ATP. Fig. 6 shows our very prelim-inary result on this reaction. In order to achieve the 50lK resolution observed here we averaged 600 framesfrom the camera and did a differential comparison be-tween the protein mixing channel and a static channelfilled with the buffer. We wish to stress that these arevery preliminary experiments that absolutely push ourtechnology in its present state.

4. Conclusions

We have discussed that a critical part of understand-ing the movement of a protein along a free energy land-

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scape as it folds (or unfolds) is measurement of the heatreleased or absorbed by the protein during that process:in other words, time-resolved thermodynamics. We havediscussed the basic statistical mechanics and thermody-namics of this process, and present a new form of diffu-sional mixer that should allow us to measure the heatrelease with time. We have presented some initial datashowing the performance of the mid-IR camera is ade-quate to obtain high resolution (0.1 mK) heat signalsin a time-resolved flow device.

Acknowledgements

This work was supported by Grants from DARPA(MDA972-00-1-0031), NIH (HG01506), NSF Nanobi-ology Technology Center (BSCECS9876771) and theState of New Jersey (NJCST 99-100-082-2042-007).Puchalla was supported by a Lewis-Thomas/BristolMyers Squibb Fellowship. Hays S. Rye of the Depart-ment of Molecular Biology, Princeton University pro-vided the GroEL/ES samples, and Daniel Adamek ofNASA Huntsville Alabama generously provided accessto the FLIR Systems camera. We thank Peter Galajdafor fabrication of the through-wafer etched structure.We acknowledge support from the Cornell UniversityNational Fabrication Center, where the deep siliconetching process of the device reported here wasperformed.

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