Dynamical aspects of TEM-1 b-Lactamase probed by molecular dynamics

DaniloRoccatanoa,d,Gianluca Sbardellab,MassimilianoAschia,GianfrancoAmicosantec,Cecilia Bossab, Alfredo Di Nolab & Fernando Mazzaa,*aDipartimento di Chimica, Ingegneria Chimica e Materiali, Universita degli Studi, V. Vetoio, 67010,L’Aquila, Italy; bDipartimento di Chimica, Universita degli Studi ‘‘La Sapienza’’, P.le A. Moro 5, 00185,Rome, Italy; cDipartimento di Scienze e Tecnologie Biomediche, Universita degli Studi, V. Vetoio, 67010,L’Aquila, Italy; dSchool of Engineering and Science, International University Bremen, Campus Ring, 1D-28725, Bremen, Germany

Received 2 February 2005; accepted in revised form 9 May 2005

� Springer 2005

Key words: domain motion, H-bonding networks, Molecular Dynamics, W-loop, TEM-1 b-lactamase,water bridges

Abstract

The dynamical aspects of the fully hydrated TEM-1 b-lactamase have been determined by a 5 ns MolecularDynamics simulation. Starting from the crystallographic coordinates, the protein shows a relaxation inwater with an overall root mean square deviation from the crystal structure increasing up to 0.17 nm,within the first nanosecond. Then a plateau is reached and the molecule fluctuates around an equilibriumconformation. The results obtained in the first nanosecond are in agreement with those of a previoussimulation (Diaz et al., J. Am. Chem. Soc., (2003) 125, 672–684). The successive equilibrium conformationin solution shows an increased mobility characterized by the following aspects. A flap-like translationalmotion anchores the W-loop to the body of the enzyme. A relevant part of the backbone dynamics implies arotational motion of one domain relative to the other. The water molecules in the active site can exchangewith different residence times. The H-bonding networks formed by the catalytic residues are frequentlyinterrupted by water molecules that could favour proton transfer reactions. An additional simulation,where the aspartyl dyad D214–D233 was considered fully deprotonated, shows that the active site isdestabilized.

Abbreviations: DynDom – dynamic domain; ED – essential dynamics; MD – Molecular Dynamics;RMSD – root mean square deviation; RMSF – root mean square fluctuation.

Introduction

Class A serine b-lactamases represent the majorsource of bacterial resistance. These enzymes con-stitute a threat to human health since they canhydrolyze the b-lactam antibiotics according to theacylation-decylation pathway [1–5]. Therefore, theyrepresent an important target for drug design [6, 7].

Although numerous crystallographic, spectro-scopic, molecular mechanics and dynamics simula-tion, and site-directed mutagenesis studies havebeenperformed (many references are reported in thereview of Matagne et al. [3]), the catalytic mecha-nism and the exact role of the active site residues arestill controversial and the relationship betweenvariants and their catalytic properties are not yetcompletely understood [3, 8].

TEM-1 (Figure 1), the prototype of TEMfamily of class A b-lactamases, is formed by

*To whom correspondence should be addressed. Fax: +39-0862-433753; E-mail: mazza@univaq.it

Journal of Computer-Aided Molecular Design (2005) 19: 329–340DOI 10.1007/s10822-005-7003-0

329

two domains, an a/b and an all a domain [9]. Thea/b domain includes a five-stranded antiparallelb-sheet onto which three helices (h1, h10 and h11)are packed. The a domain, composed by eightelices (h2–h9), is placed on the other side of theb-sheet. The two domains are connected by twohinge regions. Three salt bridges (E37–R61, R43–E64, R61–E64) and one H-bond (E37–V44) in thefirst hinge region, the salt bridge (R222–D233) andone H-bond involving the aspartyl dyad (D214–D233) in the second hinge region, prevent localstructural modification to occur. The active site isin a groove between the two domains and it issurrounded by conserved elements bearing the fiveresidues S70, K73, S130, E166, K234 moreinvolved in the catalytic process. S70 and K73are located at the beginning of h2 helix, S130 onthe so-called invariant SDN loop forming one sideof the catalytic cavity, E166 on the W loop formingthe bottom edge of the active site and K234 on theinnermost strand (s3) of the b-sheet forming theother side of the catalytic cavity.

A 1 ns Molecular Dynamics (MD) simulationof TEM-1 in aqueous solution was reported[10]. The moderate deviations and fluctuationsobtained in this simulation, with particular regardto the W loop and the complex hydrogen bondingnetworks found in the active site, reproduce the

situation found in the crystal structure of TEM-1[9].

It is known that proton transfer reactionslargely depend on dynamical factors like thepersistence of H-bonding network, the presenceand residence time of water molecules and themobility of residues. Since the dynamical behav-iour of the free enzyme is a first step for thecomprehension of the effects of the ligand bindingin the active site, we have performed a 5 ns MDsimulation of TEM-1 in aqueous solution. Startingfrom the crystallographic coordinates, the proteinshows a relaxation in water with a Root MeanSquare Deviation (RMSD) from the crystal struc-ture increasing up to 0.17 nm, within the firstnanosecond. Then a plateau is reached and themolecule fluctuates around an equilibrium confor-mation. The results obtained in the first nanosec-ond are in agreement with those of the previoussimulation [10]. The successive equilibrium con-formation of the protein in solution shows anincreased overall mobility. A relevant part of thebackbone dynamics implies a rotational motionsof one domain relative to the other. Moreover aflap-like translational motion anchores the termi-nal part of the W loop to the body of the protein.Some H-bonds, both intramolecular and withwater molecules, show a fluctuating behaviour.

Figure 1. Cartoon representation of TEM-1 b-lactamase. The relevant structural elements are indicated.

330

Moreover, different residence times characterizethe water molecules in the active site and theH-bonding networks of the catalytically relevantresidues are frequently interrupted by water mol-ecules, that could facilitate the proton transferreactions. These represent new aspects that can notbe observed in the first nanosecond of the simu-lation. Finally, the effect of the protonation stateof the aspartyl dyad D214-D233 on the enzymestability was probed by performing another MDsimulation where the same aspartyl dyad wasconsidered fully deprotonated.

Methods

The starting model for simulations was taken fromthe crystal structure of TEM-1[9] (PDB code:1BTL). The sulfate ion found in the active site wasremoved and replaced by water. The enzyme wassolvated with water in a rectangular periodic box of6.2�7.8�6.7 nm. The water molecules makingcontacts smaller than 0.15 nm with protein atomswere removed. The total number of water moleculeswas� 9000. The simple point chargemodel [11] wasused to describe the water molecules. Seven sodiumcounter-ions were added to neutralize the total boxcharge. Two different simulations, each of the samelength (5 ns), considering the D214–D233 dyad inthe mono and fully deprotonated state, respectivelywere performed. According to pKa calculations [12]the D214 was considered as the protonation site.However, similar results (data not reported) havebeen obtained in a 1 ns trial simulation consideringthe D233 as protonation site. In the simulations thetemperature was maintained at 300 K by theisothermal coupling method [13]. The LINCS algo-rithm [14] was used to constrain all bond lengths.Rotational and translation motions of the proteinwere removed using the rototraslational constraintmethod [15]. A unitary dielectric permittivity wasused. The calculation of the long range interactionswas performed by the Particle Mesh Ewald method[16], using a grid spacing of 0.12 nm combined witha fourth-order B-spline interpolation to computethe potential and forces in between grid points. Anon-bond pairlist cutoff of 0.9 nmwas used and thepairlist was updated every four steps. All atomswere given an initial velocity obtained from aMaxwellian distribution at the desired initial tem-perature. After the minimization, the system was

equilibrated by 100 ps of MD run with positionrestraints on the protein atoms. After the equilibra-tion, the position restraints were removed and thesystem was gradually heated from 50 K to 300 Kduring 50 ps. All the MD runs and the analyses oftrajectories were performed using the GROMACSsoftware package [17].

To study the changes in backbone structurealong the simulations, we have used the combinedapproach of the essential dynamics (ED) analysisand the dynamic domain (DynDom) identificationmethod proposed by Hayward [18]. This combinedapproach has been successfully employed to ana-lyze the dynamics of domains in different proteins[19, 20].

The Essential Dynamics analysis[21] allows thecharacterization of collective motions (essentialmotions) occurring during the MD simulation.The analysis was performed by building thecovariance matrix of the positional fluctuationsof the backbone atoms obtained from MD simu-lations. Upon diagonalization of the covariancematrix, a set of eigenvectors and their correspond-ing eigenvalues is generated defining a new set ofgeneralized coordinates. The eigenvectors corre-spond to directions in a 3 N dimensional space(where N is the number of atoms used for theanalysis) along which collective fluctuations ofatoms occur. The eigenvalues represent the totalmean square fluctuation of the system along thecorresponding eigenvectors. By projecting thetrajectory of the protein on the first two eigenvec-tors, the two configurations having the oppositelargest displacement along the corresponding ei-genvectors are extracted from each projections andused to perform the identification of the domainmotion using the program DynDom [22].

Results and Discussion

Deviations and fluctuations

The time course of the backbone atoms RMSD ofTEM-1 simulated in aqueous solution, relative tothe crystal structure, is reported in Figure 2. Thecurve reaches a stationary value of 0.17 (s.d. 0.01)nm after 1 ns. It should be observed that the timespent in reaching the equilibration is significantlylonger than that of the entire previous simulation(�1 ns) [10]. In order to identify the enzyme region

331

affected by major deviations, the RMSD of thebackbone atoms of each residue, computed in thelast 2 ns of the simulation, is reported in Figure 3(top panel). The bordering regions of each domainpresent large deviations and the largest one affectsthe W loop region. The bottom panel of Figure 3shows the Root Mean Squared Fluctuation(RMSF) of the backbone atoms of each residuecomputed in the last 2 ns. A comparison betweenthe top and bottom panel of Figure 3 allows todeduce that the regions with large RMSD havelow fluctuations. This indicates that the structuralmodifications of the enzyme, once in aqueoussolution, are steady in time.

Domain motion analysis

The ED analysis of the simulated trajectory showsthat, over a total of about 800 degrees of freedom,80% of the overall motion is spread over the first44 eigenvectors and the first two eigenvectorscontribute for 50% of the overall fluctuations. TheDynDom analysis reveals the presence of only onemechanical hinge axis that is described by the firsteigenvector. This mechanical axis is displaced lessthan 0.4 nm from the Ca of residues 64, 185–187,232, 246, 247 and 260, that belong to the hingeregions or are in their proximity. Consequently,the structural domains match the dynamicaldomains which are involved in a rotation ampli-tude of 12� around this axis, with 91% of closure

motion. A colour representation of the enzymeregions (red and blue) involved by the rotationalmotion togheter with the hinge axis is given inFigure 4. The five-stranded b-sheet with helices h1and h11 are the elements storing the elastic energyfor the dynamical motion of the enzyme.

Structure of the W loop in water

The RMSD of the backbone atoms reported inFigure 3 reveal that the W loop region (161–179),forming one edge of the active site, is affected bylarge deviations and the P174 is the residueshowing the largest deviation (>0.45 nm). More-over, this significant structural modification isconcerted with the formation of a new H-bondbetween the N175 N of the W loop and the R65CO of the body of the protein. As soon as thiscontact reaches H-bonding values (�0.3 nm) theRMSD adopts an average steady value (�0.2 nm).Both events take place simultaneously at thebeginning of the simulation, evolve for about500 ps and remain steady for the rest of the

Figure 3. RMSD (top) and RMSF (bottom) of the backboneatoms for each residue of TEM-1 in aqueous solution alongthe last 2 ns of simulation. The vertical lines indicate the posi-tions of the catalytic residues S70, K73, S130, E166, K234.The strands, helices and W loop regions are indicated by dot-ted, striped and full blocks. The a and a/b domains are alsoindicated.

0 1 2 3 4 5

Time (ns)

0.10

0.15

0.20

0.25

RM

SD (

nm)

Figure 2. Time course of RMSD of the backbone atoms ofTEM-1 in aqueous solution with respect to the crystal struc-ture.

332

simulation. The positions adopted by the W looprelative to the body of the protein are presented inFigure 5 by the red and blue ropes standing for theinitial (crystal) and final (water) situation, respec-tively. This flap-like translational motion, anchor-ing the W loop to the body of the protein, may beimportant for the enzyme activity, as it may helpto prevent a complete hydration of the loop inaqueous solution, maintaining the catalyticallyimportant E166 in the vicinity of the active S70residue. This motion of the W-loop is not reportedin the previous simulation [10] and differs from theone observed in the MD simulation of the PC1b-lactamase from S aureus [23]. In the last case aconformational transition of the W-loop is linkedto the rupture of the important salt bridgeN�R164-Od2D179.

The following four salt bridges, N�R164–Od2D179, N�R178–Od2D176, Ng1R164–O�1E171,N�R161–Od1D163, found in the crystal structure

of TEM-1 [9], confer polar character to the W-loopand are important for maintaining its conforma-tion. Their time evolutions during the simulationwere examined. The N�R164–Od2D179 contact,linking the two ends of the loop, maintains anaverage steady value along the entire simulation.This salt bridge is an invariant among class Ab-lactamases. The N�R178–Od2D176 contact van-ishes after about 500 ps and shows a fluctuatingbehaviour in the rest of the simulation. Thecontact Ng1R164–O�1E171 becomes increasinglylarger in the simulation. Finally, the N�R161–Od1D163 salt bridge is lost after about 700 ps and,at about 2 ns of simulation, the N�R161 engagesOH158 in a new H-bond that is maintained for therest of simulation.

Three H-bonds, OT181–NF66, NT181–OF66,NR161–OT180 were reported in the crystal struc-ture of TEM-1 [9], as further stabilizing the N andC termini of the loop. The time evolutions of the

Figure 4. Representation of motion of the red regions relative to the blue ones occuring in the MD simulation. The arrow indi-cates the hinge axis, green the regions where the hinge bending takes place and black those regions not involved in the motion.

333

first two H-bonds show steady average valuesalong the entire simulation, indicating stableH-bonding, while the maintainance of the thirdH-bond appears more perturbed along the trajec-tory.

Water bridges in the active site

The presence of water molecules H-bonded toactive residues can be significant for the catalyticactivity since it lowers the energy barrier in theproton transfer reactions involved in the acylationand deacylation of substrates. On the purpose wehave examined the H-bonding networks formed bywater molecules simultaneously engaging at leastthree H-bonds with the surrounding residues ofthe active site.

The time course of water bridged by D214,K234 and S235 residues is reported in Figure 6a. A

water is simultaneously H-bonded with theseresidues for a time longer than 3 ns. Afterwards,it is immediately replaced by a new one formingthe same network lasting until the end of thesimulation. This stable presence of water is impor-tant since the catalytic K234 residue can have atdisposal a slowly mobile water molecule. Thisnetwork neatly corresponds to that found in theTEM-1 crystal [9].

The time course of water bridged by S70, E166and N170 residues is reported in Figure 6b. Theblank spaces there reported do not stand for waterlack, but simply indicate absence of water simul-taneously forming three H-bonds. Eleven differentmolecules exchange this position in the 5 nssimulation. The longest residence time is about1 ns, indicating a water exchange faster than theprevious case. The water bridging the S70 hydro-xyl and E166 carboxylate groups has also been

Figure 5. Flap-like translational motion of the W loop. The red and blue ropes represent the position adopted in the crystal and insolution, respectively. The dotted line indicates the N175-R65 H-bond formed in solution.

334

found in the TEM-1 crystal and in other class Ab-lactamase crystal structures [9, 24–26]. It hasbeen suggested that this conserved water may actas a relay in the reaction transfer of proton from

S70 to E166 in the acylation step of the catalyticprocess. This proposal has been validated byrecent findings. Electron Nuclear Double Reso-nance (ENDOR) spectroscopy revealed the pres-ence of a water forming H-bonding with E166 inthe acylenzyme of TEM-1 [27]. Further evidenceemerges from the crystal structure of the adductbetween a boronic acid inhibitor covalently linkedto the OcS70 of the M182T TEM-1 mutant [28].The interesting result emerging from this ultrahighresolution structure is the protonated state of theE166 carboxylate group, indicating that a protoncan be shuttled from S70 to E166 through theintermediate water. Even the MD simulation ofTEM-1 [10] shows that the S70-W-E166 associa-tion is stable throughout the trajectory probed andthe water does not exchange with the bulk. Oursimulation not only confirms this result but,because of longer simulation times, it shows thepossibility of water exchange. This is in betteragreement with the functional role that the watershould play at this site.

The active S70 is also involved, with thecooperation of the other two catalytic residuesK73 and S130, in another network of H-bondsaround water molecules differing from those pre-viously examined. The time evolution is reportedin Figure 6c. This alternative interaction of S70might be considered in view of the still acylatingcapacity of the E166N TEM-1 mutant [29–31]. Inthis case the acylation should rely on an alternativemechanism without the direct participation ofE166. One proposal assigns an active kinetic roleto K73 and S130 residues that are strictly con-served in class A b-lactamases.

The presence of a water molecule in the‘‘oxyanion hole’’ was also investigated. Severalwater molecules were found to bridge the residuesS70 and A237 NH groups in the time period of thesimulation, with a maximum residence time of300 ps and a minimum below 50 ps.

H-bonding networks of catalytic residues

The time course of the contact between OcS70 andO�E166 is reported in Figure 7a. These residuespractically never form direct H-bond along thesimulated trajectory. Therefore, E166 cannotdirectly accept a proton from S70. This event canonly be mediated by a water molecule assisting thenucleophilic attack of OcS70 on the substrate

Figure 6. (a) Occurence of the water bridge with residuesK234, D214 and S235; (b) Occurence of the water bridge withresidues S70, K73, E166 and N170; (c) Occurence of the wa-ter bridge with residues K73, S70 and S130.

335

lactam carbonyl group, as discussed and shown inFigure 6b. This behaviour differs from that ob-served in the MD simulation of PC1 [23] where a

short hydrogen bond (�2.8 A) OcS70–O�E166forces the water away, indicating an S70-W-E166unstable cluster.

Figure 7. Time courses of the reported distances.

336

The NfK73 and S130 carbonyl oxygen formdirect H-bond only during the first nanosecond ofsimulation (Figure 7b). This is in accordance withthe average value of 0.29 nm found for thiscontact in the 1 ns MD simulation [10]. However,our simulation shows that after 1 ns the directH-bond is no longer maintained. In fact, this iscompetitive with the water bridging the sameresidues, as reported in Figure 6c.

The salt bridge NfK73–O�E166 distance islonger during the first nanosecond of simulationand becomes shorter in the remaining part (Fig-ure 7c). This interaction is competitive with theH-bond formation between the K73 ammoniumand S130 CO groups (Figure 7b). As long as thetwo residues are engaged in H-bonding, the saltbridge is weaker, becoming stronger once theH-bond is no longer present. It has been proposedthat an uncharged K73 might be obtained by theproton transfer K73 fi E166 in the presence ofsubstrate. In this way K73 could play the role ofgeneral base activating the OcS70 [32].

Even the H-bond NfK73-OcS70 (Figure 7d) isin competition with that between NfK73 and S130carbonyl oxygen. The close proximity of the S70and K73 side-chains, H-bonded even in the TEM-1crystal [9], has suggested the alternative hypothesisof the likely involvement of K73 as the general basein the acylation reaction [3]. According to thishypothesis the K73 side-chain would act as theproton abstractor to increase the nucleophilicity ofS70. However, the K73 terminal side-chain shouldbe deprotonated to fulfil this task. On the otherhand, it has also been observed that the positivelycharged alkylammonium group of K73 helps inorienting the S70 hydroxyl for efficient catalysis [3].

OcS130 forms direct H-bond with NfK234(Figure 7e), OcS70 (Figure 7f) and NfK73 (Fig-ure 7g), respectively. The almost simultaneousinterruption of these interactions can be correlatedto a water bridging K73 and S130 (Figure 6c). Theinteraction with K234 shows smaller fluctuationsthan those with the other two residues. Recently, ithas been proposed that the acid catalyzed proton-ation of b-lactam N is favoured over the basecatalyzed nucleophilic attack to the carbonylcarbon of lactam group [33]. According to thisproposal, the N-protonation, initiating event ofthe catalytic reaction, would be catalyzed by aH-bonded cluster including the substrate carbox-ylate group, a water molecule and the couple of

H-bonded S130 and K234 residues playing the roleof proton donor. The direct H-bond S130–K234interrupted by a water bridge, may well representthe preorganized aggregation in aqueous solutionof free enzyme before substrate intervention.

An alternative pathway, for delivering theproton from S70 to the leaving b-lactam N inthe acylation reaction, might be mediated by thesubstrate carboxylate with the following stepsS70–S130-COO)

substrate-Nsubstrate. This mechanismis similar to that of class C b-lactamases that issupported by the crystal structure of a complexformed by the S69G mutant of AmpC [34]. TheOcS70–OcS130 direct H-bond (Figure 7f) mayrepresent the step following the proton transferfrom S130 to the substrate carboxylate.

Previous MD simulations [10] have shown thatthe polar cluster K73–E166 is linked to theOcS130–NfK234 H-bond through the OS130–NfK73 interaction. Our simulation shows thatthis link is lost after 1 ns, however it can bemaintained through the other long lasting H-bondOcS130–NfK234.

Enzyme stability and protonation state of theD214–D233

A key role in stabilizing the second hinge region ofTEM-1 is played by D233. In fact, this residuesimultaneously forms a salt bridge and a H-bondwith R222 and D214, respectively. The contactOdD214–OdD233 of 0.28 nm, found in the crystal[9], indicates that one of the aspartates is likelyprotonated in the crystallization conditions(pH=7.8). Arguments favouring D214 as beingprotonated have been reported [12]. Although it isgenerally agreed that only one of the aspartates isprotonated for most of the time, the protonlocation likely changes during enzyme reaction[35, 36]. A search of the lowest contact between thecarboxylate oxygens of D214 and D233 among thecrystal structures of TEM family, at a resolution<0.2 nm, reveals that the majority of these valuesfall in the range 0.24–0.31 nm. In few cases, alwaysin presence of potassium ion, significantly largervalues have been found, the largest one being0.83 nm for TEM-32 (PDB code: 1L10). Surpris-ingly, this structure reveals that the potassium ionforms a salt bridge with OdD233 (0.27 nm) shiftingthe OdD214 to form a new short contact (0.27 nm)

337

with NgR222. This indicates that the H-bondedaspartyl dyad can be easily destabilized.

We have also investigated the influence ofprotonation state of the D214–D233 dyad on thestructural properties of TEM-1. On the purpose anadditional MD simulation was performed for theenzyme with the fully deprotonated aspartyl dyad.The RMSD reaches a stationary value of 0.22 nmsignificantly larger than that obtained in the othersimulation (Figure 2), thus indicating the effectcaused by this proton removal. In order to givedetails on the residues and interactions involved inthis structural modification, in Figure 8 we reportthe time course of three contacts monitored in thesimulation of TEM-1 with the monopronated(black curve) and fully deproronated (grey curve)aspartyl dyad, respectively. The contact OdD214–OdD233 (Figure 8a) maintains a steady valueindicating a stable H-bond (black curve) in agree-ment with the crystal structure, whereas it becomesincreasingly larger (grey curve), because of repul-sion between the negatively charged carboxylateoxygens. The salt bridge NgR222–OdD233 (Fig-ure 8b) shows a stationary value of 0.34 nm (blackcurve) in accordance with the crystal, whereas

values larger than 0.7 nm at the end of thesimulation (grey curve) severely weaken thisattractive interaction. The contact of 0.4 nmbetween OdD214 and NfK234 formed after 2 ns(grey curve of Figure 8c) is indicative of a new saltbridge that cannot be present with the monoprot-onated dyad (black curve) because of the proton-ated D214. Therefore, the fully deprotonatedaspartyl dyad destabilizes the interactions con-necting the second hinge region of TEM-1. Inaddition (data not shown) the deviations andfluctuations of the active site residues from thecrystal structure, are larger than those obtained inthe simulation of TEM-1 with the monoprotonat-ed dyad. It is interesting to observe that theinvariant D233, present in all class A b-lactamases,forms a salt bridge and a H-bond, both stabilizingthe enzyme second hinge region. Consistently, ithas been reported that replacement of D233 byother residues, strongly reduces the TEM-1 cata-lytic activity [37]. The H-bond OdD214–OdD233and salt bridge NgR222–OdD233 connecting thesecond hinge region are destabilized both in D233TEM-1 variants and in TEM-1 with the fullydeprotonated aspartyl dyad.

Figure 8. Time course of the indicated contacts monitored in the simulation of TEM-1 with the monopronated (black curve) andfully deproronated (grey curve) aspartyl dyad, respectively.

338

Conclusions

A 5 ns trajectory of the TEM-1 b-lactamase inaqueous solution has been obtained by MD sim-ulation. As shown in figure 2, the RMSD, withrespect to the crystal structure, increases in the firstnanosecond up to a plateau of 0.17 nm, then anequilibrium conformation is reached, that fluctu-ates around an average structure. The resultsobtained in the first non equilibrated part do notsignificantly differ from the ones previously re-ported by Diaz et al. [10]. Differently, in theequilibrated part of the simulation an increasedmobility is observed. The dynamical behaviour ischaracterized by a closure motion around an axispassing near the two hinge regions. As a conse-quence the two structural domains pratically matchthe dynamical ones. The W loop, forming one edgeof the catalytic region, undergoes a flap liketranslational motion anchoring its terminal partto the body of the protein through the new H-bondNN175–OR65. The water bridge S70-W-E166 is along-lived interaction, and our simulation revealsthat this conserved water can also exchange withthe bulk on a ns time scale, emphasizing itsfunctional role. Some H-bonds, both intramolecu-lar and with water molecules, show a fluctuatingbehaviour. The complex H-bonding network foundin the active site among the catalytically importantresidues, often interrupted by water molecules, canbe relevant for the catalytic activity. Several preor-ganized H-bonded clusters proposed for differentproton transfer reactions can be recognized. Thisgreat variety of possibilities can give reason that theroute for acylation cannot be unique and alterna-tive ways might be possible. The MD simulation ofTEM-1 with the fully deprotonated aspartyl dyadD214–D233 shows that the R222–D233 salt bridgeand the D214–D233 H-bond, important interac-tions connecting the second hinge region, aredestabilized. A similar destabilization should occurin the practically inactive D233 TEM-1 variants.MD simulations of TEM-1 complexed with sub-strates are being currently performed.

Acknowledgements

This work was supported by grant from MIUR(PRIN 2003 on Structure and dynamics ofredox protein to A. D. N.). The authors thank

Dr. S. Hayward, University of East Anglia, for usefuldiscussions and support of DynDom program,Prof. S Mobashery, Univerity of Notre Dame,Indiana (U.S.A) for his criticism. CASPUR-Romeis acknowledged for computational facilities.

References

1. Waley, S.G. Page, M.I. (Ed.), The Chemistry of b-lactams1992, pp. 198–228, Chapman and Hall, Glasgow, U.K.

2. Galleni, M., Lamotte-Brasseur, J., Raquet, X., Dubus, A.,Monnaie, D., Knox, J.R. and Frere, J.-M., Biochem.Pharmacol., 49 (1995) 1171.

3. Matagne, A., Lamotte-Brasseur, J. and Frere, J.-M., Bio-chem. J., 330 (1998) 581.

4. Wright, G.D., Chem. Biol., 7 (2000) 127.5. Walsh, C., Nature, 406 (2000) 775.6. Strynadka, N.C., Martin, R., Jensen, S.E., Gold, M. and

Jones, J.B., Nat. Struct. Biol., 3 (1996) 688.7. Ness, S., Martin, R., Kindler, A.M., Paetzel, M., Gold, M.,

Jensen, S.E., Jones, J.B. and Strynadka, N.C., Biochemis-try, 39 (2000) 5312.

8. Yang, Y., Rasmussen, B.A. and Shlaes, D.M., Pharmacol.Therap., 83 (1999) 141.

9. Jelsch, C., Mourey, L., Masson, J.M. and Samama, J.-P.,Proteins, 16 (1993) 364.

10. Diaz, N., Sordo, T.L., Merz, K.M. Jr. and Suarez, D.,J. Am. Chem. Soc., 125 (2003) 672.

11. Berendsen, H.J.C., Grigera, J.R. and Straatsma, T.P.,J. Phys. Chem., 91 (1987) 6269.

12. Swaren, P., Maveyraud, L., Guillet, V., Masson, J.-M.,Mourey, L. and Samama, J.-P., Structure, 3 (1995) 603.

13. Allen, M.P. and Tildesly, D.J. Computer Simulation ofLiquids. Oxford University Press, Oxford, 1987.

14. Hess, B., Bekker, H., Berendsen, H.J.C. and Fraaije,J.G.E.M., J. Comput. Chem., 18 (1997) 1463.

15. Amadei, A., Chillemi, G., Ceruso, M.A., Grottesi, A. andDi Nola, A., J. Chem. Phys., 112 (2000) 9.

16. Darden, T.A., York, D.M. and Pedersen, L.G., J. Chem.Phys., 98 (1993) 10089.

17. van der Spoel, D., van Drunen, R. and Berendsen, H.J.C.,GROningen MAchine for Chemical Simulation, Depart-ment of Biophysical Chemistry, BIOSON Research Insti-tute, Nijenborgh 4 NL-9717 AG Groningen, 1994.

18. Hayward, S. and Berendsen, H.J.C., Proteins, 30 (1998)144.

19. deGroot, B.L., Hayward, S., vanAalten, D.M.F., Amadei,A. and Berendsen, H.J.C., Proteins, 31 (1998) 116.

20. Daidone, I., Roccatano, D. and Hayward, S., J. Mol. Biol.,339 (2004) 515.

21. Amadei, A., Linssen, A.B.M. and Berendsen, H.J.C., Pro-teins, 17 (1993) 412.

22. Hayward, S. and Lee, R.A., J. Mol. Graph. Model., 21(2002) 181.

23. Vijayakumar, S., Ravishanker, G., Pratt, R.F. and Beve-ridge, D.L., J. Am. Chem. Soc., 117 (1995) 1722.

24. Knox, J.R. and Moews, P.C., J. Mol. Biol., 220 (1991) 435.25. Lamotte-Brasseur, J., Dive, G., Dideberg, O., Charlier, P.,

Frere, J.M. and Ghuysen, J.M., Biochem. J., 279 (1991)213.

26. Herzberg, O., J. Mol. Biol., 217 (1991) 701.

339

27. Mustafi, D., Sosa-Peinado, A. and Makinen, M.W., Bio-chemistry, 40 (2001) 2397.

28. Minasov, G., Wang, X. and Shoichet, B.K., J. Am. ChemSoc., 124 (2002) 5333.

29. Strynadka, N.C., Adachi, H., Jensen, S.E., Johns, K.,Sielecki, A., Betzel, C., Sutoh, K. and James, M.N.G.,Nature, 359 (1992) 700.

30. Guillaume, G., Vanhove, M., Lamotte-Brasseur, J.,Ledent, P., Jamin, M., Joris, B. and Frere, J.-M., J. Biol.Chem., 272 (1997) 5438.

31. Massova, I. and Mobashery, S., Antimicrob. Agents Che-mother., 42 (1998) 1.

32. Lietz, E.J., Truher, H., Kahn, D., Hokenson, M.J. andFink, A.L., Biochemistry, 39 (2000) 4971.

33. Atanasov, B.P., Mustafi, D. and Makinen, M.W., Proc.Natl. Acad. Sci. U.S.A., 97 (2000) 3160.

34. Beadle, B.M., Trehan, I., Focia, P.J. and Shoichet, B.K.,Structure, 10 (2002) 413.

35. Kashparov, I.V., Popov, M.E. and Popov, E.M., Adv. Exp.Med. Biol., 436 (1998) 115.

36. Torshin, I.Y., Harrison, R.W. and Weber, I.T., ProteinEng., 16 (2003) 201.

37. Denisov, V.P. and Halle, B., Faraday Discuss., 103 (1996)227.

340