Allosteric effect of 2′-adenylic acid on the Pseudomonas pyridine nucleotide transhydrogenase

J. ikfol. Biol. (1972) 70, 651-664

Allosteric Effect of 21-Adenylic Acid on the Pseudomonas Pyridine Nucleotide Transhydrogenase DANIELD.LOTJIE-/',NATHAITO.KAPLANAXDJAMES

D. MULEAN$

Departments of Chemistry and Biology University of Cdifirnia, 8an Diego La Jolla, Calif. 92037, U.X.A. (Received 7 June 1971, and in revised form 12 iWay 1972) In highly purifkd Pseudomonas pyridine nucleotide transhydrogenase, allosteric a&iv&ion by 2’-adenylic acid is eooompanied by changes of reactivity and sedimentation resulting from alterations in protein conformation. The requirement for 2’-sdenylic acid is specific for the activation of the enzyme-catalyzed oxidation of DPNH by TPN+. Direct effect of the 2’-adenylia acid on the dissociation of the protein is demonstrated by a decrease in the sediment&ion coefficients, from S 20,w = 110 s to a lower SzO,W= 28 s. A shift in the rate of attainment of equilibrium is observed as a result of the 2’-adenylio aoid activation. High-resolution electron microscopic studies reveal the structural details of the protein. The enzyme exists as an unbranohed Lament of 500 A to 5000 A in length and 80 A to 100 A in width. The polymerio forms appear to be helical in structure. Activation by 2’-edenylic acid induces a dissociation of the fllamentous form into cyliuclrical subunits of 100 B to 200 ,& in diameter. Each unit appears to consist of 6 to 8 lobes regularly arranged at the periphery. These results are discussed in relation to the chemical and physical characteristics of this macromolecular system.

1. Introduction Bacterial pyridine been demonstrated (Colowick, Kaplan,

nucleotide transhydrogenase from Pseudomonas aeruginosa has to catalyze the following reaction without additional substrate Neufeld & Ciotti, 1952; Kaplan, Colowick & Neufeld, 1952) : TPNH

+ DPN+

--t TPN+

+ DPNH.

The transhydrogenase readily catalyzes the reduction of DPN+ by TPNII, but the oxidation of DPNH by TPN+ proceeds at a relatively insignificant rate. In addition, Kaplan et al. (1952) reported that the reactions in which TPN+ is the product do not proceed to equilibrium witbin a reasonable time. This phenomenon has been attributed to the fact that TPN+ is an extremely effective inhibitor of the enzyme. Subsequent studies by Kaplan, Colowick, Neufeld & Ciotti (1953) demonstrated that 2’-adenylic acid is capable of stimulating the oxidation of DPNII by TPN+ . Because a highly purified enzyme was obtained, Cohen & Kaplan (1970a) were able to ascertain t Present address: Pro&or end Gamble Co., P.O. Box 39175, Cincinnati, Ohio 45239, U.S.A. $ Present address: Division of Chemical Physics, C.S.R.I.O., P.O. Box 160, Clayton, Victoria, Au&r&a. 651

65.2

D.

that the 2’adenylic

D. LOUIE,

N.

0.

KAPLAN

acid is capable of altering

AND

J. D. MCLEAN

the sedimentation

characteristics

of the

enzyme under conditions of enzyme activation. Similar activation effects by the 2’-adenylic acid have been demonstrated on the transhydrogenase from Azotobacter vilzelandii (Louie & Kaplan, 1970a; Chung, 1970), but without effect on the animal mitochondrial transhydrogenase (Kaplan, Colowick & Neufeld, 1953). However, 2’-adenylio acid has been found to be a competitive inhibitor of the Chromutium transhydrogenase (Keister & Hemmes, 1966) and a number of TPN-linked dehydrogenase systems (Neufeld, Kaplan & Colowick, 1955). Our studies attempted to resolve whether the 2’-adenylic acid, which has a 2’phosphate group similar to that of the TPN + , functions directly as an antagonist of the inhibition of TPN+ or as an allosteric effector of the enzyme. In the studies presented in this paper, data from electron micrographs are shown which demonstrate the allosteric effect of the 2’-adenylic acid on the conformational transitions of the Pseudomonaa transhydrogenase.

2. Materials and Methods (a) Enzyme source and a;ssay The pyridine nucleotide transhydrogenase was purified from and assayed as described previously (Cohen & Kaplan, 1970a; One unit of enzyme was defined as the reduction of 1 pmole of reduced coenzyme. The reaction rate was monitored at A,,, nm at A400 l’Un = 11.3 X lo3 M-l.

Pseudomonas aerz@nosa Louie & Kaplan, 19’70b). (TN)DPNt per min by a and the Ed of (TN)DPNII

(b) Analyt&zl centrifwattin Measurements were carried out on a Spinco model E analytical centrifuge equipped with a photoelectric scanning system. Sedimentation studies were followed by the increase in absorption of the (TN)DPNH at AaoO nm, catalyzed by the enzyme sedimenting under standard assay conditions (Cohen t Kaplan, 1970a). A type II center piece (Beckman SP no. 350259) for cells (12 Mm, 4-O” single sector) was used. All the reagents for the assay experiment were prepared in a O-1 M-Tris.HCl buffer (PH 7-5) containing O-15 M-Nacl. The enzyme was layered onto the cell through a small well situated between the top sapphire window and the reaction medium, which was connected by a narrow cha,nnel. After acceleration of the rotor reached the necessary speed (5000 to 8000 rev./mm), the enzyme was then forced into the medium in the sector, thereby initiating the reaction. All experiments were carried out at 27°C. (c) Radioactivity transfer studies Synthesis of [B] [4-3H]TPNH and @] [4-3H]DPNH were achieved as previously (TPN described (Louie & Kaplan, 19705) and the preparation of the [7-14C]nicotinamide and [7-14C]nicotinamide) DPN was according to the procedure of Everse, Kaplan & Schiohor (1970). Radioactivity wasmeasured with a Beckman liquid-scintillation counter, using 10 ml. of Bray’s (1960) scintillation fluid. All radioactive starting compounds were purchased from the New England Nuclear Corporation. Antisera were prepared in rabbits against purified Pseudomonas aemginosa transhydrogenase. The course of immunization was started with a 2-mg sample of the enzyme in Freunds complete adjuvant injected intra-musoularly for 4 consecutive weeks and then followed by a booster of 1 mg protein sample after an interval of 2 months. The rabbits were bled through the marginal ear vein one month after the initial injection and one week after each booster. All antisera were stored at - 10°C without purification. 3-(AcPy)TPN+, oxidized t Abbreviations used: (!CN)DPN+, oxidized thionicotinamid+DPN; 3-scetyl-pyridine-TPN; 2’P-ADPR, 2’-phosphoedenoside diphosphate ribose; ~&O-AMP, (t-isoadenosine)-W-phosphate.

PSEUDOMONAS

653

TRANSHYDROGENASE

Micro-complement fixation was carried out according to the procedure of Wasserman & Levine (1961). The reaction buffer mixture contained 0.14 M-NaClj0.01 M-Tris.HCl/ (pH 7.45). Reaction 0.05 mi?r-MgSO,/O~15 mu-C&1,/0*1 y. b ovine serum albumin (Armour) times were from 14 to 20 hr at 4°C. (e)

Fluorescence and

polarization

of jhorescerxe

Fluorescence measurements were performed on a Zeiss Spectrofluorometer model ZFM4C. The excitation wavelength for the flavin fluorescence was at 365 nm. Polarization of flavin fluorescence spectra were obtained on a polarization spectrofluorometer designed by Weber & Bablouzian (1966). The polarization of fluorescence, P, was defined as (111- 1,/J,, +I,), with Iii and 11 representing the intensities parallel and perpendicular to the polarized excitation. A pair of Corning C53-72 filters was used to separate the emission from the excitation radiation during the measurement of the polerization of FAD fluorescence. (f) Electron vnicroscopy Specimen grids of copper coated with fenestrated Formvar films, covered with an ultrathin carbon film were prepared by the method developed by Fern&ndez-Mor&n, van Bruggen & Ohtsuki (1966). The transhydrogenase was diffused onto the specimen grid as a monolayer. A small amount of finely divided talc was sprinkled evenly over the surface of the 0.1 m-potassium phosphate buffer (pH 7.5) containing 10 mM-2-mercaptoethanol and 1 mM-EDTA. A droplet of transhydrogenase solution was gently placed on the buffer surface with a fine, freshly drawn glass pipette. The leading edge of the spreading protein monolayer pushed back the talc, leaving a clear circular area. The prepared grid was held with forceps and touched on to the monolayer surface, thus allowing the protein to diffuse onto the carboncovered grid. After a period of 1 to 2 min, the excess solution floated off on a piece of filter paper without fixation. Specimens were stained by floating on the 1% uranyl formate or phosphotungstic acid (pH 6.6) for 2 to 5 min. Finally, the specimen was allowed to air-dry after blotting the edge of the grid onto e, piece of filter paper. Examinations were carried out on a Phillips 300 electron microscope with an operating voltage at 80 kV. Photographs were taken with Kodak high-contrast photographic plates.

(g) Stopped-Jlow studies Stopped-flow experiments were carried out on the Aminco-Morrow stopped-flow appar&us (American Instrument Co., Inc.). Buffer system contained 0.1 m-potassium phosphate with 10 mM-2-mercaptoethanol. All experiments were carried out at 25°C.

3. Results (8) Effects of activators

on oxidation

of

.DPNE

It is of importance to note that reactions in which TPNH is the oxidant do not require activators (Kaplan, Colowick & Neufeld, 1953; Cohen & Kaplan, 1970a,b). A 2’-phosphate derivative appears to be essential for transhydrogenase activity in reactions involving DPNH as the oxidant?. Table 1 demonstrates the influence of the various mononucleotides on the DPNH oxidation. It is evident that 2’-adenylic acid is an effective activator, the presence of 2’-adenylic acid stimulating the rate of oxidation of DPNH by TPN+ some 60-fold. Other monophosphates of adenosine, including the 2’,3’-cyclicadenylic acid, were found to be totally ineffective as an activator of the DPNH/TPN+ reaction. Similarly, 2’-CMP, 2’-GMP and 2’-UMP exhibited absolutely no activating effect at concentrations comparable to the 2’-AMP. Although 3-(AcPy)TPN+ is not a normal substitute in the transhydrogenase reactions, it is relatively t It is possible to follow this reaction beclause the maximum absorption of TPNH whereas that of the reduced thionicotinamide analogue (!CN)DPNH is at 400 urn.

is at 340 nm,

654

D. D.

LOGIE,

X.

0.

KAPLAN TABLE

AND

J. D.

MCLEAN

1

Effects of azi%iva.!omon oxidation of DPNH DPNH f TPN+ + TPNH + DPN+ Nuoleotide None 2’-AMP 3’-AMP V-AMP 2’,3’-cyclic AMP 3’,6’-ayolio AMP 2’-CMP 2’-GMP 2’-UMP 3-(AcPy)TPN+ 2’P-ADPR t&o-AMP

do.D./min 0.004 0.240 0.005 0.004 0.010 0.004 0.004 0.003 0.002 0.175 0.100 0.004 0.001

Each ass4~y medium of 1.0 ml. contained 0.3 mu-DPNH; 0.6 mar-(TN)DPN; buffer (pH 7.5); 0.1 m-nucleotide; Pseudomonas transhydrogenase, 0.6 unit.

100 mna-Tris. HCl

effective MI a substitute activator for the oxidation of DPSH reaction. In addition, 2’P-ADPR shows some activating power even though ADP and ATP completely lack the capacity of stimulating bacterial transhydrogenase reactions. NXN+ , which is a poor reactant, also shows little or no activating effect. (b) Effeectof 2’-adenylic acid on dissociatim of tramhydrqenme Previous studies by Cohen & Kaplan (197Oa) demonstrated that the 2’-AMP and TPX+ are capable of altering the sedimentation characteristics of the enzyme. At high protein concentrations the enzyme sediments as a single homogeneous component only in the proscnce of the 2’-adenylic acid as monitored by scanning the flavin absorption. Questions ha.vc been raised as to whether the 2’-AMP does, in fact, kinetically activate the reactions of the enzyme (Cohen & Kaplan, 1970a) simultaneously with dissociation, or whether t.he dissociation process at high protein concentrations is independent of t,he 2’-adenylic acid effect. l!his problem was resolved by st,udying the association4issociation properties of low enzyme concentrations. Hence an examination of the sedimentation charactsristics of the enzyme at the assay conditions was carried out. The results indicated t.hat dissociation at low protein concentration were related to the kinetic activation of the 2’-adenylic acid. Table 2 provides evidence that the presence of 2’-AMP can indeed change the sedimentation characteristics of the enzyme. In the presence of the 2’-AMP, the S20,w of 28 s is consistent with the S,,,, value obtained for the effect at high protein concentration (Cohen & Kaplan, 1970a). A variation of the band-forming sedimentation technique suggested by Vinograd, Brunner, Kent & Weigle (1963) was used in these experiments. In order to monitor the TPNH/(TN)DPN+ reaction, the photoelectric scanner W~ZJset up to follow continuously the formation of (TX)DPNH by following A400 nm, which is the absorption maximum of the reduced analoguet. Because the enzyme was 7 We have referred to the over-all reaction ae essentially irreversible. This does not mean that there is no oxidation of DPNH by TPN in the absence of 2’-AMP, but that the rate of oxidation of DPhXH is 80 slow that it msy ba insignikant kinet&Ily. A detailed disoussion of this point is given elsewhere (Kaplan, 1972).

PSEUDOMONAS

656

TRANSHYDROGENASE

TABLE 2 Effect of 2’-adenylic acid on enzyme sedimentation as measured by enzyme activity and jiavin absorption Enzymic

Addition

A B C

-2’~AMP +2’-AMP +2’-AMP

TPNH TPNH DPNH

+ (TN)DPN+ + (TN)DPN+ + (TN)DPN+

reaction

-+ TPN+ --f TPN+ --t DPN+

Activity + (TN)DPNH + (TN)DPNH + (TN)DPNH

110 30 28

S20.w Flavin 121 32 33

Centrifugation was carried out in 100 mM-potassium phosphate buffer (pH 7.5); 10 mEn-Cleland’s reagent (dithiothreitol); 1 mi%r-EDTA and 1 mm-2’-AMP where applicable. Photoelectrio scans were carried out at 440 mn for FAD scan (protein concentration 8 mg/ml.). Activity scan was determined by measuring the (TN)DPNH gamed at 400 nm (0.05 mg protein/ml.) Experiments on enzyme activity scans were run in 0.15 iu-NaCh See text for description of method. All reagents and reactants were placed in the cell, and the enzyme was layered onto the cell just before sedimentation.

sedimented into the denser solvent containing the assay components, a narrow band was formed as the result of the enzyme-catalyzed reduction of (TN)DPN. Utilizing this reduction of the (TN)DPN with either TPNH or DPNH as a measure of the enzymic activity, the catalytic activation and sedimentation characteristically couId be correlated. A concentration gradient was generated continuously by the diffusion of the enzyme to catalyze reduction of (TN)DPN+ through the solvent medium. The oxidation of TPNH catalyzed by the transhydrogenase does not require the activator, 2’-adenylic acid. An SZO,wvalue of 110 s (Table 2) was obtained for this reaction representing the sedimentation characteristics of the enzyme in the absence of 2’-adenylic acid. The S,,,, value of 110 s correlated reasonably well with the previous S20,wof 120 s, calculated for the principle sedimenting component from scanning of the flavin absorption of the enzyme at high protein concentrations (Cohen & Kaplan, 1970a). We have not ruled out the possibility that at high concentrations the enzyme can self associate. The S2,,,wvalue of 28 s (Table 2) for the enzyme-catalyzed oxidation of the DPNH in the presence of 2’-adenylic acid was consistent with the sedimentation studies utilizing high protein coefficient (AS,,,, = 33 S) from earlier ultracentrifugation concentrations (Cohen & Kaplan, 1970a). The scanner studies with low enzyme concentrations under assay conditions in the presence of 2’-AMP indicate a direct relationship between the activation characteristics and the dissociation induced by 2’-adenylic acid. The reaction rates became equalized by adjustment of the enzyme concentration, so that the rates of reduction of (TN)DPN+ were similar in the above sedimentation experiments . Xince these experiments were carried out under similar buffered conditions (pH 7-5), there should be no effect as the result of pH changes. The fact that a higher centrifugal force was necessary for the sedimentation of the 2’-AMP-activating DPNH/(TN)DPN+ reaction (31,300 rev./min) over that for the TPNH/(TN)DPN+ reaction (25,000 rev./min) further supports that there is a difference in the size in which these two reactions occur. No attempts, however, were made to account for possible differences in the diffusion coefficients or the frictional coefficients of the two enzyme conformations. 43

656

D. D. LOUIE,

N.

0.

KAPLAN TABLE

EJffectof 2’-AMP Pyridine

‘“C A:

Experiment DPNH TPN+ DPN+ TPNH

B:

Experiment Experiment

J. D. MCLEAN

3

on hydrogen transfer under equilibrium conditions

nucleotides

Experiment TPNH DPN+ TPN+ DPNH

AND

Specifia ectivity No T-AMP 3H

(cts/miu/~mole) Added 2’-AMP 14C

(lo-3

M) =H

660 -

50,000 -

300 310

24,000 100 8000

700 -

46,000 -

450 400

23,000 35,000

A [C-“H]TPNH; B [4-sH]DPNH;

[7-14C]DPN+ [7-W]TPN+

; TPN+ ; DPNH. ; DPNf ; TPNH.

(c) Radioactivity transfer studies The regulatory effect of the 2’-adenylic acid has been demonstrated by its ability to catalyze the stereospecific transfer of the hydrogen between the reduced and oxidized pyridine nucleotides (Louie & Kaplan, 1970b). Using stereospecifically titrated DPNH at the 4B position of the niootinamide ring, the enzyme catalyzes only in the presence of 2’-AMP, a direct hydrogen transfer from the 4B position of the DPNH to the 4B position of reduced TPN. In the absence of 2’-adenylic acid, no transfer of hydrogen was detected. Kaplan, Colowick & Neufeld (1953) have demonstrated that the 2’-AMP increases the rate of transfer between the TPNH and the DPN+ as catalyzed by the Pseudomonas $uorescens transhydrogenase and also increases the activation of the reverse reaction. It is of interest to note that an equilibrium constant of approximately unity was obtained from stoichiometric measurements for all the components of the reaction (Kaplan et al., 1952). The reaction appears to stop when the concentration of the reduced and oxidized forms of each pyridine coenzyme are equal. Because of the effect of 2’-AMP on DPNH oxidation by TPN + , it was of interest to determine the effect of the nuoleotide on the trsnshydrogenase reactions with the components initially at equilibrium concentrations. Results of such studies are given in Table 3. Experiments were carried out in which the pyridine coenzymes were introduced in approximately equal concentrations (6 PM). Stereospecifically labeled [B] [4-“H]TPNH and [14C]DPN + were used in the study of the TPNH oxidation by DPN + . Similarly [B] [4-3H]DPNH and [14C]TPN + were used in following the reverse reaction, oxidation of DPN + . Figure 1 shows a typical elution profile of the reaction products from the DEAE-cellulose column (Louie 82;Kaplan, 1970b); in this case, the reaction was the result of the oxidation of TPNH by DPN+, in the presence of the transhydrogenase and 2’-AMP. Tritium was transferred to the 14C-labeled DPN + from the [B] [4-3H]TPNH to form a doubly-labeled DPNH. It is interesting to note that the relative specific activity is lower than that expected for complete transfer. No tritium is found on the oxidized [l%]DPN+ because only the 4B hydrogen is being transferred to and from the [B] [4-3H]TPNH. When 2’-AMP is omitted under the above reaction

PSJUDOMONAB

657

TRANSHYDROQENASE

TPNH

600

24 !

00 d 0 04

0

0

20

40

60

00

100 120 hction

160

140

150

200

no

FIU. 1. Cbromstograpbic profiles of the pyridine nuoleotides on a DEAE-cellulose aolumn. Coenzymes were eluted with a N&l gmdient from 0 to 2% in a 6 mx-Tris-HCl buffer (pH 7-5). Optical density wes monitored et 260 n.m (-- l -- l --) for the oxidized form and 340 11111

(. . x *. x * e) for the reduced aoenzymes. Radioactivity was measured for W (-,0-O-) 3H (--n-A-) by liquid scintillation counting. AU separations wem carried out at 4°C.

and

conditions, no tritium is detected on the [14C]DPNH. Similar results were obtained when the reverse direction was followed with the following labeled compounds, [B]mixture. [4-3H]DPNH and [l*C]TPN + , as two labeled compounds in the equilibrium These results suggest that at equilibrium the enzyme is inactive and that under these conditions the enzyme becomes active only in the presence of 2’-AMP. The significance of the quantitative difference between experiments (3) and (4) is not clear at the present time. (d) Complement sxation The ultracentrifugation studies indicate that the 2’-A.&P could dissociate the enzyme into smaller sedimentation units and that conformational changes may result from the dissociation. Such conformational changes can be detected by the microcomplement fixation (C’F) technique of Levine (1962)) Van Vunakis & Levine (1963) and Reich& Bucoi, Wyman, Antonini & Rossi-Fanelli (1965). A highly specific antibody directed toward the purified Pseudomonas transhydrogenase was obtained from rabbit antisera. Because these experiments use antibody at a very low concentration (l/60,000 dilutions), only the antigens of highest affiuity will be reactivated. Figure 2 shows the results of the C-fixation experiments. A maximum Gxation of 60% was obtained for the enzyme at O-01 pg protein concentration. In the presence of 2’-AMP, the (?-fixation curve shows a defkite vertical shift (decrease) from the untreated enzyme at about the same concentration. These results indicate further that the 2’-nuoleotide is promoting a structural change. (e) Fluorescence and polarization

of flwwescence

Effects of the 2’-adenyhc acid binding on the conformational alteration of the enzyme were followed by changes in flavin fluorescence as well as of polarization of

658

D. D.

LOUIE,

N.

0.

KAPLAN

I o-~-~--~-o, L-*-J

AND

J. D.

MCLEAN

I I

,L; Tmshydrcqenase

FIQ. 2. Effect of 2’-adenyhc acid on the C-fixing of Pseudomonas transhydrogenase. The microomplement fixation by Pseudomonas transhydrogenase (-O--()-) and 2’.AMP-treated crleyme --A-A-) reacting with 8 l/60,000 dilution of anti-rabbit transhydrogenase.

FIG. 3. Flavin 5uorescence emission spectra of effects of activators on Pseudomonas transhydrogenase. Concentrations of the nuclootidcs wero at 1 mn, in a 10 mm-potassium phosphate buffer (pH 7.5) containing 1 mr,r-EDTA, and 10 mnr-jl-mercaptocthanol. Excitation wavelength was set rat 365 nm. Spectra were taken at 27°C.

fluorescence. Figure 3 illustrates the fluorescence emission spectra of the flavoprotein in excitation at 365 nm. The fluorescence of the enzyme-bound FAD exhibik little 620 nm fluorescence because of quenching; t,hus, the quenching of the FAD when bound to protein is consistent with the results observed with ma,ny ot.her flavoproteins (Massey & Williams, 1965). As demonstrated by the spectrum shown in Figure 3, the binding of 2’-AMP (low3 M) results in a fourfold enhancement of the flak fluorescence. A

PSEUDOMONAS

I O”450

659

TRANSHYDROGENASE

I 400

1 450

:

Excitation wavelength (nm)

FIG. 4. Polarization of &win fluorescence spectra of Pseudomonas transhydrogenase. Excitation spectra of the Pseudomonas transhydrogenase (--O-O-) and 1 mm-2’-AMP-treated transhydrogenase (-~----a-) were measured in 10 mM-potassium phosphate buffer (pH 7.5) containing 10 mM-fi-mercaptoethanol, and 1 m&r-EDTA, in a-2.5 ml. vol. All spectra, were taken at 27°C.

mixture of equal quantities of 2’- and 3’-AMP at 1 mM induces a similar increase in fluorescence yield. Although 3’-AMP was not an allosteric effector, it did demonstrate a capacity to bind to the enzyme as shown by a slight enhancement of flavin fluorescence. Little change was obtained in observed results with 5’-, 3’,5’-cyclic and the 2’,3’- cyclic AMP. Results in Table 1 demonstrate that the 3-(AcPy)TPN+ and the 2’-P-ADPR are relatively good activators of the transhydrogenase reactions. These two compounds exhibit an enhancement of more than twofold over that of fluorescence as compared to the untreated transhydrogenase. It is of interest to note that Tl?N+ addition results in approximately a twofold increase of the flavin fluorescence. Possible conformational changes of the enzyme were measured with polarization of flavin fluorescence over a range of excitation wavelengths. Figure 4 shows that the native enzyme exhibited a maximum fluorescence polarization of 0.38 as indicated by the open-circle spectrum. After treatment with 2’-AMP (1 mM), the polarization of fluorescence of the flavoprotein had increased to a polarization maximum of 0.42. This is indicated by the spectrum with the triangles. The increase appears to be significant and the polarization is comparable to other flavoproteins (Table 4). (f) Electron microscopy The Pseudomonas transhydrogenase dissociation into smaller functional units by 2’-AMP is indicated by the complement fixation and sedimentation results. Electron microscopy studies, primarily involved with the negative-staining technique on a monolayer, produced most convincing evidence to support the 2’-AMP effect on the enzyme’s dissociation characteristics. The Pseudomonas transhydrogenase has an unbranched filamentous structure of non-uniform length in the range of 500 A to 5000 A (Plate I). However, predominant filamentous rods measuring between 500 A and 200 A in length comprised the majority of the population. Under high-contrast negative-staining conditions, these structures appear to consist of regular crossstriations with indentations along the filament axis, suggestive of a helical structure (Plate II). One can observe the presence of two distinctly different images (Plate III).

660

D.

D. LOUIE,

N.

0.

KAPLAN

AND

J. D. MCLEAN

TABLET Maximum jiuorescence polarization values of $avin enzymes and Jlavin nucleotides Enzyme

or coenzyme

P8eudonzonas transhydrogemse + T-AMP Lipoyl dehydrogenase Glut&Gone reductase D-Amino acid ox&we FAD FMN

Polarization,

P

0.38 0.42 0.41 0.20 0.065 0.033 0.015

The widths of the majority of the filamentous helix have an apparent dimension of 80 A to 100 A. In stereoscopic views these structures appear to be loosely coiled into an open helix with a pitch of about 100 A to 200 A along the longitudal axis of the fZament. A second image frequently observed is shown to have an apparent dimension of 90 A to 100 A square. Scattered among these images are some circular segments of similar sizes. After treatment of the enzyme with 2’-AMP, the total population is transformed into an exclusively smaller unit (Plate IV). The circular projection of the particle is about 120 A to 150 A in diameter. Regular rectangular images (100 A to 120 A) are assumed to be on the side view of the circular space. If this supposition is true, the general morphology of the 2’-AMP-treated particle is cylindrical in shape. The view of the “subunit” structure (Plate V) indicates 6 or 8 units peripheral to the particle, projecting some 30 A to 35 A out from the core, which may possibly be hollow, with the protein forming the outer spiral coat. Assuming a nominal density for a cylinder of protein of 100 A high and 120 A in diameter, the molecular weight would be in the range of 900,000. The Svedberg value for a protein of such size and shape should be approximately 20 to 25 S; this is consistent with the SzO,Wvalues obtained from the hydrodynamic data as discussed above. Based on the value of one FAD cofactor per 45,000 gram of protein (Cohen & Kaplan, 197Oa), each resulting structure from 2’-adenylic acid activation is composed of about 20 subunits of 40,000 to 45,000 molecular weight. (g) XtoppdjTOw .a&23 Sedimentation and electron-microscopic studies have demonstrated that the 2’-AMP activation is correlated with the dissociation of the transhydrogenase. Kgure 5 summarizes stopped-flow experiments carried out on the enzyme in order to determine the time factor involved in the activation by 2’-AMP. Reactions were carried out under assay conditions, and the course of the reaction was followed at 400 nm, which is the absorption maximum of the reduced (TN)DPN. The reactions were initiated by the rapid mixing of the contents from the two syringes. When the enzyme was preincubated with 2’-AMP in one syringe, no lag period was detected. However, when the enzyme was placed in one syringe and the 2’-AMP, DPNH and (TN)DPN+ were put in the second syringe, a lag period of 100 to 150 milliseconds was observed on a rapid mixing of the two syringes. The only experimental conditions under which DPNH is the substrate, and a lag is not observed, is when the enzyme is preincubated

PLATE I. Electron miorograph of a typical preparation of native transhydrogenase enzyme, negatively stained with 1 oj uranyl formate.

Pseudomonas x 144,000.

aeruqinosa

Lfaeingp.

G60

PLATE II. Electron Details of the helical

micrograph of an isolated nature of the filamentous

portion protein

of the negatively are illustrated.

stained transhydrogenase. x 314,300.

PLATE III. stained

with

Micrograph of a representative 1 o/0 many1 formate to demonstrate

transhydrogenase enzyme in the untreated state the existence of the subunit structures. x 308,000.

PLATE 2’adenylic

IV. Electron micrograph acid. x 257,100.

of a preparation

of transhydrogenase

after

treatment

with

PIXTE V. Micrograph of the enlarged subunits of 2’-AMP-treated show the mosaic arrangement of the lobes. x 535,700.

enzyme.

Subunit

structures

PSEUDOMONAS

TRANSHYDROGENASE

661

Time hsec)

FIG. 5. Stopped-flow experiments on the 2’-AMP effect on the initial reaction rate of DPNH oxidation. Reactions were carried out under assay conditions (see Materials and Methods) with the enzyme in one syringe and the reaotion mixture in the second syringe. The curve (-e--a-) represents the initial rate of DPNH oxidation when the enzyme is preincubated with the 2’-AMP (1 IIIM) before mixing. Curve (-A-A-) represents the rate of reaction when the enzyme is not inoubated with the 2’-AMP before initiation of the reaction.

with the 2’-AMP. No lag periods were produced by the 2’-AMP reactions involving TPNH. It is apparent that reaotions involving oxidation of DPNH catalyzed by the enzyme are dependent on the activation ensuing from the binding of 2’AMP.

4. Discussion The data presented above indicate that 2’-adenylic acid functions as an allosteric effector of the Pseudomonas transhydrogenase. Kaplan, Colowick & Neufeld (1953) have suggested that the 2’-AMP activation may possibly be due to the competitive action of the 2’-phosphoadenosine for the TPN +. By competing with the TPN+ in a manner similar to that observed with a number of TPN-linked dehydrogenases (Neufeld et al., 1955) the activation by the 2’-AMP could allow for the oxidation of the DPNH. Kinetic data also support such a possibility (Cohen & Kaplan, 1970b). The results indicate that the 2’-phosphoryl group and the adenine moiety are of primary importance in promoting the allosteric effect. Substitution of the 2’-phosphate group at either the 3’- or 5’hydroxyl group of the ribosyl moiety abolishes the allosterio effect. Replacement of the adenine by other purine and pyrimidine bases results in the same inactivity. Because the 2’,3’-cyclic AMP gives little activation, one can postulate that the 2’-phosphate group must be a monoester. The sedimentation, complement fixation, fluorescence and electron-microscopic studies all point to the concept that 2’-AMP induces a conformational change, which is associated with dissociation of the very large complex into the smaller units. The stimulation by 2’-AMP of DPNH oxidation may be the result of dissociation of the protein thus exposing new binding sites. The bacterial transhydrogenase represents an interesting example of the role the enzyme plays in regulating the rate at which equilibrium of a reaction is attained. The oxidation of TPNH by DPN+ is catalyzed by the enzyme, but the corresponding oxidation of DPNII by TPN+ requires the activation of 2’~AMP. The attainment of

662

D.

D. LOUIE,

N.

0.

KAPLAN

AND

J. D. MCLEAN

this equilibrium is governed by the enzyme through its effective control of the rates in opposite directions. The animal transhydrogenase freely catalyzes reversibly the oxidation and reduction of the pyridine nucleotides (Kaplan, Colowick, Neufeld $ Ciotti, 1953). The 2’-AMP offers no stimulation to the animal transhydrogenase. However, the reaction rates in the two directions as catalyzed by the animal enz”yme are not equal (Stein, Kaplan & Ciotti, 1959; Kaufman & Kaplan, 1961). It appears that the 2’-AMP causes a shift in the rate of the bacterial transhydrogenase reaction. Experiments with stereospecifically labeled pyridine nucleotides have (at equimolar concentration) indicated that there is no exchange of labels when the reaction is carried out in the absence of 2’-AMP. The addition of 2’-AMP catalyzes a shift in the equilibrium and thus allows the reaction to become activated. It appears that there is a formation of new enzyme species enabling such an exchange reaction to occur. This is similar to the allosteric control of 5’-AMP on the biodegradive n-threonine dehydrogenase of Escherichia coli thereby causing activation accompanied by a shift in the equilibrium between the monomer and the tetramer forms (Phillips & Wood, 1965 ; Whanger et al., 1968). Hydrodynamic studies of the Pseudomonas enzyme at catalytic levels demonstrate that the protein dissociates into smaller components from the allosteric effect of 2’-AMP. It appears that the 2’-AMP must rapidly activate the dissociation of the enzyme into the active conformer so that the oxidation of DPNH by TPN is able to proceed. It is not possible at this time to ascertain whether this is truly a one-step process or two inter-dependent rapid sequential steps. The electron microscopy studies clearly illustrate the allosteric effect produced by 2’-AMP in dissociating the bacterial enzyme into discrete subunits. The filamentous form of the transhydrogenase is unique and the polymeric species of the enzyme have been demonstrated in the acetyl CoA carboxylase from animal tissues (Kleinschmidt, Moss & Lane, 1969) and the glutaminase systems (Kvamme, Tveit & Svenneby, 1965). Unlike the polymeric form of acetyl CoA carboxylase, the filamentous form of the bacterial transhydrogenase is the active species for TPNH oxidation by DPN. However, in the reverse reaction DPNH oxidation by TPN necessitates the dissociation of the enzyme into subunits. The resulting subunits appear to consist of a more closely packed molecular structure as observed by electron-microscopic scans. The dissociated structures of the native transhydrogenase resulting from the action of 2’-AMF show some similarities to the glutamine synthetase from Escherichia wli. Electron microscopy studies of the purified glutamine synthetase from E. coli indicate that the enzyme is a symmetrical aggregate of 12 identical subunits (Valentine, Shapiro & Stadtman, 1968). The cylindrical appearance of the Pseudomonas transhydrogenase subunits could conceivably be a face-to-face association of the units leading to a long hexagonal tubular aggregate similar to that observed for the glutamine synthetase. Unlike the glutamine synthetase, electron microscopy studies of the Pseudomonas transhydrogenase show no indication of equilibrium mixture of the native and subunit population, as a result of 2’-AMP activation. An increase in the polarization of flavin fluorescence following the addition of 2’-AMP suggests that the enzyme-bound FAD becomes less mobile. In the caseof lipoyl dehydrogenase, Setlow (1968) suggested that the FAD is held tightly in a protective “pocket” and can not rotate fully independent of the protein moiety. This decrease in rotational depolarization assumes that the lifetime of the FAD does not change upon binding to the protein. In the case of the Pseudomonas enzyme, the 2’-AMP appears to induce a type of conformational alteration so that further restriction of the FAD

PSEUDOMONAS

TRANSHYDROGENASE

663

rotation within the protein structure occurs. In addition, the dissociation of the enzyme by 2’-AMP might increase the effective size of the fluorescent molecule, FAD, in a highly restricted hydrophobic environment. In effect, the 2’-AMP probably converts the transhydrogenase from the “relaxed” filamentous form into a more “taut” species which is comparable to the glutamine synthetase system (Valentine et d., 1968). Attempts can be made to interpret the observations of the allosteric effects of the 2’-AMP. The “two-state” model of allosteric transition of Monod, Wyman & Changeux (1965) was postulated to account for an allosteric protein to comprise a small number of identical subunits and equilibrates between a small number of conformational states. Among these states, the more stable and predominant are those in which all the subunits of the protein have the same conformation. In the “induced fit” model of Koshland (1963), the conformational change is induced by and coincides with ligand binding. The large enhancement of flavin fluorescence, caused by the 2’-AMP, attests to the strong allosteric effect on the transhydrogenase upon binding of the mononucleotide. As yet we have found no equilibrium nor “hybrid” species in the allosterio protein population resulting from 2’-AMP induction; however, we have not made an extensive study of the effects of subsaturating levels of the 2’-AMP on the structure observable in the electron microscope or in the ultracentrifuge. The changes of reactivity and sedimentation resulting from the changes in the conformation of the protein nevertheless appear to be the consequences of the direct effect of ligand, 2’-adenylic acid, binding to the protein. This work was supported in part by grant no. CA0361.1 from the Institute, National Institutes of Health, and the American Cancer Society One of us (D.D.L.) was a postdoctoral fellow (no. 7FZAM31, 613-01) Institute of Arthritis and Metabolic Disease, during t#he course of these

National Cancer grant no. P-77L. of the National studies.

REFERENCES Bray, G. A. (1960). Analyt. Rio&em. 1, 279. Chung, A. E. (1970). J. Bact. 102, 438. Cohen, P. T. & Kaplan, N. 0. (19700). J. Biol. Chern. 245, 2825. Cohen, P. T. L Kaplan, N. 0. (1970b). J. Biol. Chem. 245, 4666. Colowick, S. P., Kaplan, N. O., Neufeld, E.F. & Ciotti, M. M. (1952). J. Biol. Ohem. 195,95. Everse, J., Kaplan, N. 0. & Schichor, S. (1970). Arch. Biochem. Biophy.s 136, 106. Fernandez-Mor&n, H., van Bruggen, E. F. J. & Ohtsuki, M. (1966). J. Mol. BioE. 16, 191. Kaplan, N. 0. (1972). Harvey Lecture, series 66, p. 105. New York: Academic Press. Kaplan, N. O., Colowick, S. P. & Neufeld, E. F. (1952). J. Biol. Chem. 195, 107. Kaplan, N. O., Colowick, S. P. & Neufeld, E. F. (1953). J. BioZ. Chew. 205, 1. Kaplan, N. O., Colowick, S. P., Neufeld, E. F. & Ciotti, M. M. (1953). J. Biol. Chem. 205,17. Kaufman, B. T. & Kaplan, N. 0. (1961). J. BioZ. Chem. 236, 2133. Keister, D. L. & Hemmes, R. B. (1966). J. BioZ. Chem. 241, 2820. Kleinschmidt, A. K., Moss, J. & Lane, M. D. (1969). Science, 166, 1276. Koshland, D. (1963). Cold Spr. Harb. Syrrq. Qwxmt. BioZ. 28, 473. Kvamme, E., Tveit, B. & Svenneby, G. (1965). Biochem. Biophys. Re.s. Comm. 20, 566. Levine, L. (1962). Fed. Proc. 21, 711. Louie, D. D. & Kaplan, N. 0. (1970a). Proc. of the Symp. on Pyridine Nwleotide-Dependent Dehydrogesusss, ed. by H. Sund, p. 351. New York: Springer-Verlag.

Louie, D. D. & Kaplan, N. 0. (1970b). J. BioZ. Chem. 245, 5691. Massey, V. & Williams, C. H. (1965). J. BioZ. Chem. 240, 4470. Monod, J., Wyman, J. & Changeux, J. P. (1965). J. Mol. BioZ. 12, 88. Neufeld, E. F., Kaplan, N. 0. & Colowick, S. P. (1955). Biochim. biophys.

Acta,

17, 525.

664

D. D. LOUIE,

Phillips, Reichlk,

N.

0.

KAPLAN

AND

J. D. MCLEAN

A. T. & Wood, W. A. (1965). J. Biol. Chem. 240, 4703. M., Bucci, E., Wyman, J., Antonini, E. & Rossi-Fan&i,

A. (1965). J. .MoZ. BkZ.

11, 775. Setlow, P. (1968). Thesis, Brandeis University, Waltham, Mass. Stein, A. M., Kaplan, N. 0. & Ciotti, M. M. (1959). J. BioZ. Chem. 234, 979. Valentine, R. C., Shapiro, B. M. & Stadtman, E. R. (1968). Biochemistry, 7, 2143. Van Vunakis, H. & Levine, L. (1963). Ann. N.Y. Accld. Sci. 103, 735. Vinograd, J., Bnmner, R., Kent, R. & Weigle, J. (1963). Proc. Nat. Acad. Sci., Wash. 49, 902. Wasserman, E. & Levine, L. (1961). J. Immunol. 87, 290. Weber, G. & Bablouzian, B. (1966). J. BioZ. Chem. 241, 2258. Whanger, P. D., Phillips, A. T., Rabinowitz, K. W., Piperno, J. R., Shsda, J. D. & Wood, W. A. (1968). J. BioZ. Chem. 243, 167.