X-factor (hemin) and V-factor (NAD). While the requirement for NAD was established in 1937 (4), investigation into the pyridine nucleotide metabolism of the.
JOURNALOF BIOLOGICAL CHEMISTRY 0 1986 by The American Society of Biological Chemists, Inc. THE
Vol. 261, No. 13, Issue of May 5, pp. 6016-6025,1986 Printed in U.S.A.
Characterization of Haemophilus influenzaeNucleotide Pyrophosphatase AN ENZYME OF CRITICAL IMPORTANCE FOR GROWTH OF THE ORGANISM* (Received for publication, October 7, 1985)
David W. Kahn and Bruce M. Anderson$ From the Department of Biochemistry and Nutrition, Virginia Polytechnic Institute and StateUniversity, Blacksburg, Virginia24061
A nucleotide pyrophosphatase isolated from Haernophilus influenzae was purified to electrophoretic homogeneity and characterized with respect to molecular weight, substrate specificity, pH profile, thermal stability, functional group involvement, and effectiveness of selective inhibition. The enzyme catalyzes the hydrolysis of NAD toNMN and AMP and appears located appropriately to facilitate the internalization of NAD needed to satisfythe V-factor growth requirement of the organism. In the processing of NAD and structurally related substrates, the enzyme exhibited negative cooperativity. Structural alterations in the purine moiety of these dinucleotide substrates had pronounced effects on the negative cooperativity of the enzyme. AMP, ADP, and several related nucleotides were observed to be effective substrate-competitive inhibitors of the enzyme. Several of the dinucleotides serving as substrates for the nucleotide pyrophosphatase were evaluated with respect to substituting for NAD in supporting growth of the organism. AMP and ADP inhibited growth of the organism when NAD served as V-factor, and this inhibition correlated well with the inhibitory effects of these nucleotides on the purified nucleotide pyrophosphatase.
Haemophilus influenzae typeb is the primary cause of bacterial meningitis in humans,responsible for 10,000-20,000 cases annually in the United States (1).Other members of the genus are responsible for numerous diseases in man and other animals. Treatment of H. influenzae infections has relied predominantly upon the use of antibiotics, particularly ampicillin; however, in 1974 the first reportof strains resistant to ampicillin appeared in the literature (2) and in 1978 a report estimated that 16% of all infections were resistant to ampicillin treatment (3).Haemophilus organisms are subclassified by their unique requirement for one or both of two growth factors, viz. X-factor (hemin) and V-factor (NAD). While the requirement for NAD was established in 1937 (4), investigation into the pyridine nucleotide metabolism of the organism has been minimal. It has been established that H. influenzae is incapable ofNAD biosynthesis from typical precursors of the de novo biosynthetic pathways as well as from nicotinamide and nicotinic acid (4-6). In addition, it is * These studies were supported by Research Grant DMB8508930 from the National Science Foundation. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. 3 To whom correspondence should be addressed.
known that both nicotinamide riboside and nicotinamide mononucleotide (NMN) arecapable of serving as V-factor (68), suggesting the presence of a salvage pathway or pyridine nucleotide cycle. The noted inability of the highly polar NAD molecule to diffuse across various biological membranes suggests the possibility that thefunctioning of NAD as V-factor for H. influenzae would require the hydrolysis of NAD to a smaller, transportable fragment. Preliminary studies of this process (9) demonstrated the presence of a nucleotide pyrophosphatase in this organism that catalyzes the degradation of NAD to NMN and AMP. The present study reports the purification and characterization of this enzyme and an investigation into the manipulation of the growth of the organism by selective interactions with this enzyme and the resulting perturbation of the pyridine nucleotide metabolism of this organism. EXPERIMENTALPROCEDURES~ RESULTS
Demonstration of Nucleotide Pyrophosphatase ActivityWashed cells of H. influenzae were incubated with NAD and the extracellular mediumwas analyzed for nucleotides by reverse phase high performance liquid chromatography. As shown in Fig. 1, the extracellular NAD was rapidly degraded within 20 min to NMN and adenosine. Adenosine was produced by the combined action of the nucleotide pyrophosphatase and either 5’-nucleotidase or alkaline phosphatase activity. Using the appropriate extinction coefficients, stoichiometric quantities of adenosine were recoveredin the extracellular medium. At 20 min, however, only 65% of the predicted amount of NMN was present and, unlike adenosine, which remained at a constant concentrationfrom 20-40 min, NMN dropped to 48% of the value predicted for total hydrolysis of the NAD. Nicotinamide and ADP-ribose were not observed at any time in the extracellular medium, indicating the absence of NAD glycohydrolase activity. Purification of the Nucleotide Pyrophosphatase-The enzymewas produced ina soluble form for purification by conversion of the cells to spheroplasts using lysozyme and EDTA. Release of the enzyme required the presence of both Portions of this paper (including “Experimental Procedures,” part of “Results,” Figs. 1S-8S, and Table IS) are presented in miniprint at the endof this paper. Miniprint is easily read with the aid of a standard magnifying glass. Full size photocopies are available from the Journal of Biological Chemistry, 9650 Rockville Pike, Bethesda, MD 20814. Request Document No. 85M-3356, cite the authors, and include a check or money order for $6.40 per set of photocopies. Full size photocopies are also included in the microfilm edition of the Journal thatis available from Waverly Press.
6016
H. influenzae Nucleotide Pyrophosphatase 0
6017
minutes
IO minutes
1 r
ADENOSINE
20 minutes
L
,NMN
f
n
o
E
N
o
s
l
N
E .IO
40minutes
.20
.30 .40 .50 RS
NMN ADENOSINE
FIG.1. High performance liquid chromatography analysis of the hydrolysis of NAD by cells of H. influenzue. Incubation mixtures were prepared as described under “Experimental Procedures.”
TABLE I Purification of the H. influenzae nucleotide mv’ophosphatase Fraction
Total Total proactivity tein
m 1. Lysozyme digest 731.4 2. Ammonium sulfate 282.9 3. Phosphocellulose 15.5 4. Matrex Green Gel A 1.0 5. Matrex Blue Gel A 0.3
units
3823 1814 1694 1007 928
Specific activity
unitslmg
Yield Purification %
5 100 6 47 109 44 1007 26 3500 700 24
-fold
1 20 201
lysozyme and EDTA, and with varied EDTA concentrations paralleled the release of the periplasmic enzyme 2’,3’-cyclic phosphodiesterase (14). The nucleotide pyrophosphatase was purified as shown in TableI, using ammonium sulfate precipitation, ion-exchange and affinity chromatography procedures described in the Miniprint. The enzyme was reproducibly purified to electrophoretic homogeneity, 700-fold relative to the supernatant fraction following lysozymedigestion, with a recovery of 24% of the initial activity. Properties of the Purified Enzyme-The molecular weight of the enzyme was determined under native and denaturing conditions by gel-filtration chromatography and sodium dodecyl sulfate gel electrophoresis, respectively. As shown in Fig. 2, the apparent molecular weights were M, = 62,500 and 65,800, respectively. The purified enzyme was shown to consist of 16%carbohydrate by weight, andthe amino acid composition was determined (see Miniprint). The UV-visible absorption spectrum consisted of a single maximum at 275 nm. The c:& was equal to 47.8. The fluorescence spectrum of the enzyme revealed excitation and emission maxima at 286 and 337 nm, respectively, indicative of the presence of tryptophan. The purified enzyme was thermolabile, and the rates of thermal denaturation, aswell as theeffects of temperature on catalytic activity, are documented in the Miniprint. The rate of hydrolysis of NAD, as catalyzed by the nucleotide pyrophosphatase, was directly proportional to theamount of enzyme present. Investigation into the effects of various cations on activity revealed that theenzyme was only slightly more active in the presence of increasing concentrations of
.30
15
.45
.60
.75
.90
KD
FIG. 2. Molecular weight determination of the nucleotide pyroph0sphatase.A shows gel filtration on Sephacryl S-200 column equilibrated with 50 mM Tris-HC1, pH 8.0, 200 mMKC1. Molecular weight standards were ( I ) yeast alcohol dehydrogenase, (2)horse liver alcohol dehydrogenase, (3) bovine serum albumin, ( 4 ) a-chymotrypsinogen A. B shows sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Molecular weight standards were ( I ) yeast alcohol dehydrogenase, (3) bovine serum albumin, (4)a-chymotrypsinogen A, ( 5 ) rabbit muscle phosphorylase a, (6) glucose oxidase, (7) heavy chain, humanIgG, and (8)ovalbumin. The arrows in A and B indicate the determined positions of the H. influenzae nucleotide pyrophosphatase.
40
00
120
160
200
FIG.3. The effect of NAD concentration on the initial velocity of the nucleotide pyrophosphatase-catalyzed hydrolysis of NAD. Initial velocities were determined using the titrimetric assay described under “Experimental Procedures.” Reactions contained 50 mM KC1,2 pg of enzyme, and theindicated amounts of NAD at 37 “C in a total volume of 3.0 ml.
6018
Pyrophosphatase Nucleotide H. influenzae TABLE I1 Substrate smcificitv. Low
Substrate
Nicotinamide 5.5 adenine dinucleotide 72.6 2.7 3-Acetylpyridine adenine dinucleotide 3-Aminopyridine 2.3adenine dinucleotide 127.4 1.6 Pyridine 3.6 adenine dinucleotide 35.7 1.7 Nicotinic acid adenine dinucleotide 1,4-Dihydronicotinamide 2.4 adenine dinucleotide 32.2 1.5 Flavin 4.2 adenine dinucleotide 67.8 2.3 Nicotinamide hypoxanthine 3.3 dinucleotide 114.6 1.7 Nicotinamide guanine dinucleotide 3.7 Nicotinamide 1,N6-ethenoadeninedinucleotide 3.6 3-Aminopyridine l,N6-ethenoadenine dinucleotide Nicotinamide mononucleotide" Adenosine 5'-monophosphate" Adenosine 5'-diphosphate" Adenosine diphosphoribose 32.4 4.7 Uridine diphosphoglucose 49.8 2.0 bis-p-nitrophenylphosphate"
6.3
High
K,
V"l
K"l
V,
PM
unitsf mg
PM
unitsfmg
5.4 6.2 56.1 3.2 7.9 10.9 20.0 14.8 25.5 632.0 1290
1.3 1.9
22.3
3.65.1
39.8
2.2
9.4 5.9
4.6
~~
Did not serve as substrate at 1 mM.
potassium, sodium, and magnesium ions. Relative to these rates, rates observed with increasing concentrations of calcium ions were slightly lower. Enzyme activity was not affected by the presence of 5 mM EDTA. The effect of pH was investigated over the range from pH 6.0 to 9.0. Activity was maximal from pH 8.0 to 9.0, and decreased rapidly at lower pH. Substrate Specificity-The purified nucleotide pyrophosphatase catalyzed the hydrolysis of a variety of dinucleotides and structurally related compounds with relatively the same degree of efficiency. The enzyme, in utilizing all substrates except several modified in the adenine ring, functioned in a manner indicative of negative cooperativity. As typified by NAD studies (Fig. 3), double-reciprocal plots of initial velocities of NAD hydrolysis were biphasic and concave downward. Extrapolation to x- and y-axis intercepts of these plots allowed for calculation of kinetic constants for the hydrolysis of all substrates, as shown in Table 11. Changes in the substituents of the pyridine ring, e.g. substitution of an isoalloxazine ring for pyridine (FAD) and eliminationof the second heterocyclic base altogether (ADP-ribose, UDP-Glc) affected the ability of the enzyme to utilize these compounds only modestly. The purified enzyme did not show phosphatase or phosphodiesterase activity. Substitution of other purine bases for adenine altered the manner in which the enzyme functioned with these substrates. Whereas substitution of hypoxanthine for adenine resulted in less favorable kinetic constants relative to NAD, substitution of guanine for adenine eliminated the biphasic kinetics observed with the otherdinucleotides. Using substrates containing ethenoadenine, such asc-NAD2 (Fig.4) and c-AAD,altered the functioning of the enzyme extensively. Both ethenoadenine-containing compounds analyzed were acted upon by the enzyme in a non-cooperative manner and displayed higher Michaelis constants compared to their adenine-containing parent compounds, NAD and AAD. Hill plots of the enzyme-catalyzed hydrolysis of all substrates confirmed the apparent negative cooperativity in the functioning of the enzyme and allowed for the qualitative assessment of the ability of these compounds to induce negThe abbreviations used are: €-NAD,nicotinamide 1,"ethenoadenine dinucleotide; AAD, 3-aminopyridine adenine dinucleotide; cAAD, 3-aminopyridine 1,M-ethenoadenine dinucleotide; HPLC, high performance liquid chromatography; Nir, nicotinamide riboside.
.I 4
c
0
-I
1
2
3
'
4
5
6
7
8
I
Fj FIG. 4. The effect of e-NAD concentration on the rate of hydrolysis of the dinucleotide. Reaction mixtures were prepared as described in the legend to Fig. 3.
ative cooperativity. Hill coefficients, obtained from the slopes of these plots, decrease with increasing negative cooperativity. Ainslie et al. (18) have proposed that the ratio of high to low maximal velocity values may also serve as an indicator of the degree of negative cooperativity, wherein higher values are indicative of greater cooperativity. Hill coefficients, K,, and V, ratios for all substrates areshown in Table 111. Hydrolysis of NAD was catalyzed by the nucleotide pyrophosphatase with the greatest degree of negative cooperativity of all the substrates analyzed, and substrates that did not display biphasic kinetics had Hill coefficients approximately equal to one. Inhibition of Enzyme Activity-On addition of 20 pM AMP to reaction mixtures, the biphasic kinetics observed in the hydrolysis ofNAD were abolished and simple hyperbolic kinetics were observed (Fig. 5). The Hill coefficient for the hydrolysis of NAD in the presence of 20 p~ AMP was 0.87. The inhibitor constants for AMP and other inhibitors are shown in Table IV. The affinity for binding of adenine compounds increased in theorder ADP > AMP > adenosine. The affinity of the enzyme for NMN was significantly less than its affinity for the other half of NAD, AMP.
6019
H. influenzae Nucleotide Pyrophosphatase TABLEI11 Negative cooperativity in the functioningof the nucleotide pyrophosphatase Substrate
Nicotinamide adenine dinucleotide 13.4 Pyridine adenine dinucleotide 11.1 Uridine diphosphoglucose 8.4 Nicotinamide hypoxanthine dinucleotide 7.7 Nicotinic acid adenine dinucleotide 5.0 3-Acetylpyridine adenine dinucleotide 3.6 Flavin adenine dinucleotide 3.4 Adenosine diphosphoribose 3.0 1,4-Dihydronicotinamide adenine 3.0 dinucleotide 3-Aminopyridine adenine dinucleotide0.931.4 2.3 Nicotinamide guanine dinucleotide Nicotinamide 1,N6-ethenoadenine dinucleotide 3-Aminopyridine 1,N6-ethenoadenine dinucleotide
,500 I
I
I
I
I
2.0 2.1 2.3 2.2 1.4 1.5 1.8 1.6 1.6
0.26 0.49 0.61 0.61 0.51 0.42 0.62 0.69 0.73 0.91 0.91 1.12
I
1
I 1
.400
Demonstration of NAD Pyrophosphorylase Activity-Since NMN is produced from NAD by nucleotide pyrophosphatase activity and can substitute for external NAD for growth of the organism, it was of interest to investigate the intracellular reformation ofNAD from NMN and ATP as catalyzed by NAD pyrophosphorylase. A soluble fraction, obtained from cells of H. influenzae as described under “Experimental Procedures,” was observed to catalyze the synthesis of NAD from NMN at a rate of 71.7 nmol/min/ml of sonicate. Growth Studies-As a complement to theinvestigations of the H. influenzae nucleotide pyrophosphatase, experiments were conducted to investigate the ability of numerous compounds to either 1)serve as V-factor or 2) inhibit growth of the organism. The growth of the organism, measured turbidimetrically, with various concentrations of NAD is shown in Fig. 6. Concentrations of NAD >0.1 pg/ml readily supported growth. Doubling times for all growth studies were estimated from the time needed for the optical density at 660 nm to increase from a value of0.2-0.4. The organism also grew readily with NMN as V-factor, although at equivalent concentrations, growth with NAD as V-factor was moreefficient, as shown in Table V. The growth rates obtained at saturating NMN were belowthose obtainedat maximal NAD concentrations. Several NAD analogs which served as substrates for the nucleotide pyrophosphatase were tested for their ability
\
.-cE
TABLEIV Inhibition of the nucleotide pyrophosphatase
,300
\
Inhibitor
.200
Ki fiM
Adenosine” Adenosine 5”monophosphate Adenosine 5”diphosphate Adenosine 5”sulfate Guanosine 5‘-monophosphate Nicotinamide mononucleotide“
.IO0
0
20
40
60
80
1 0 0
120
140
No inhibition observed a t 20 p FIG. 5. The effectof AMP on the rates of hydrolysis of NAD. Reaction mixtures were prepared as described in the legend to Fig. 3. Line 1 and line 2 are reactions in the absence and presence of 20 pM AMP, respectively.
~
15.1 1.6 51.6 21.9
.
2.2 I .B 0
Fluorescence Studies-The addition of adenosine deriva- 0’ID I.4 tives to the enzyme resulted in the quenching of the fluores- 0 I.o cence of the enzyme in a concentration-dependent process. When plotted as a double-reciprocal plot, the data obtained 0.6 with adenosine, AMP, and ADP were biphasic, indicative of at least two different modes of binding of these compounds 0.2 to theenzyme (see Miniprint). Dissociation constants extrap50 1 0 0 1 5 0 200 250 300 350 400 450 500 olated from these data correlated well with the inhibitor constants determined from kinetic data. TIME (min) Effect of NAD on the Apparent Molecular Weight of the FIG. 6. Growth of H. influenzae with NAD as V-factor. The Enzyme-The effect ofNAD on theapparent molecular procedure for growth of the organism was as described under “Experweight of the enzyme was determined using gel-filtration high imental Procedures.” performance liquid chromatography. In the absence of NAD, TABLEV the nucleotide pyrophosphatase exhibited a molecular weight T h e ability of NMN and NAD to serve as V-factor of 67,800, consistent with the values obtained with both Doubling times Sephacryl S-200 column chromatography and sodium dodecyl Concentration sulfate gel electrophoresis. When 100 or 400 p~ NAD was NMN NAD present in the elution buffer, the enzyme exhibited apparent min d m l molecular weight of 104,800 and 108,100, respectively. 206.3 0.01 94.1 Chemical Modification of the Purified Enzyme-The62.5puri0.03 61.5 0.1 57.9 49.1 fied nucleotide pyrophosphatase was inactivated by 2,3-bu0.3 42.3 from these tanedione and by Woodward’sReagent K. Results56.1 1.0 53.8 38.6 studies are described and presented in the Miniprint.
H. influenzae Pyrophosphatase Nucleotide
6020
TABLEVI The ability of various compounds to serve as V-factor Each compound was present a t a concentration of 1.0 pg/ml. Doubling Compound time
TABLE VI1 Inhibition of the growth of H. inflfluenzae with NAD as V-factor All inhibitors were present at a concentration of 10.0 d m l . Doubling Compound time
min
NAD NMN 3-Acetylpyridine adenine dinucleotide Nicotinamide hypoxanthine dinucleotide NADH Thionicotinamide adenine dinucleotide Nicotinamide 1,N6-ethenoadeninedinucleotide 2.0
I
I
I
I
38.6 53.8 266.7 42.4 44.3 225.8 406.2
min
49.1 95.7 NG" 69.9 NG" 88.6 97.4 56.6 156.0
None AMP ADP ADP-ribose 3-Aminopyridine adenine dinucleotide Pyridine adenine dinucleotide 3-Methylpyridine adenine dinucleotide 4-Aminopyridine adenine dinucleotide 3-Aminopyridine 1,N6-ethenoadeninedinucleotide
I a
NG, no growth observed.
p q AAD 1 m
TABLEVI11 The effect of AMP on the growth of H.influenzae with NMN o r NAD o r V-factor Doubling times
I .6
I .2
Q
V-factor concentration
0
0
0.8
NAD No AMP
dml 0.4
n ".-7
0
100
200
52.6 51.9 300
0.003 140.0 184.2 0.01 NG" 57.6 62.7 0.0372.0 56.3 60.0 49.1 0.10 54.1 48.7 42.3 0.30 400 500 49.6 53.5 1.0 49.8
TIME (min)
FIG. 7. Inhibition of growth of H. influenzae by AAD with NAD as V-factor. The growth inhibition experimental procedures are as described under "Experimental Procedures."
to serve as V-factor. A comparison of the doubling times produced with several compounds present at a concentration of 1.0 pg/mlis shown in Table VI. Whereas the hypoxanthine analog of NAD was comparable to NAD in itsability to serve as V-factor, the other analogs were clearly less efficient. At 10 pg/ml, all the analogs produced doubling times that were still 15 min or longer than the39 min doubling time observed for NAD at 1.0 pg/ml. Several compounds were analyzed for their ability to inhibit the growth of H. influenzae in the presence of NAD. Fig. 7 shows the inhibition of growth of the organism by AAD. AAD was one of the most effective inhibitors, completely inhibiting growth at 1pg/ml. Other compounds studied as growth inhibitors are listed in TableVII. Adenine nucleotides were effective as growth inhibitors, with ADP being more effective than AMP. Although AMP effectively inhibited growth with NAD as V-factor, it did not inhibit growth when NMN was used as V-factor (Table VIII). DISCUSSION
H. influenzae, and several other members of the genus Haemophilus, exhibit a requirement for NAD as a growth factor. This growth requirement is entirely unique to this genus. Most microorganisms acquire pyridine nucleotides by either de m u 0 biosynthetic pathways or by use of pyridine nucleotide cycles (19). H. influenzae does not possess the ability to synthesize NAD de m u o , and the ability of the organism to utilize NMN as V-factor, documented here and in other studies (4-6), implies the existence of a functional pyridine nucleotide cycle. The predominant pyridine nucleotide cycles in prokaryotes, the four-membered PNC IV, and
NMN
3.6 pglml
No
AMP AMP
10.0pglml AMP
min
143.5 94.1 61.5
38.6 NG, no growth observed.
the six-membered PNC VI, are both initiated by cleavage of NAD at thepyrophosphate bond. In this study, an externally directed nucleotide pyrophosphatase was purified and characterized. The enzymewas found to be a glycoprotein released from the cell by lysozymeEDTA digestion. Successful solubilization by this procedure and the observed parallel release of the periplasmic 2',3'cyclic phosphodiesterase suggest a periplasmic location for the nucleotide pyrophosphatase. The specific activity of the purified enzyme in catalyzing the hydrolysis of NAD was 5.5 pmol/ml/mg of protein. The purified nucleotide pyrophosphatases from bovine seminal fluid (20), yeast (21), and Proteus vulgaris (22) had reported specific activities of 2.2, 0.09, and 3.7 pmol/min/mg, respectively. The purified nucleotide pyrophosphatase was found to exist as asingle polypeptide chain with an M , of 64,100. The yeast enzyme also consists of a single polypeptide with a molecular weight of 65,000 (21), whereas those obtained from mammalian sources are considerably larger (23). Unlike several bacterial nucleotide pyrophosphatases which are heat-activated (22, 24), the H. influenzae enzyme was thermolabile and was readily denatured at temperatures exceeding 15 "C. Although the optimal pH range of 8-9 is typical of most nucleotide pyrophosphatases (20,21, 25-28), the lack of apparent metalrequirements and insensitivity to EDTA is unusual (29-31); however, there are known exceptions, such asthe bovine seminal fluid (20) and ratliver lysozomal (32) nucleotide pyrophosphatases. Most nucleotide pyrophosphatases have broad substrate specificities (20-22, 25-29, 33) and the H. influenzae enzyme also displayed this property. With the exception ofAAD, structural analogs of NAD which possessed various modifications of the pyridine ring were hydrolyzed with relatively the same degree of efficiency. FADreadily served as substrate for the enzyme, as did compounds which lacked a second
H. influenzae Nucleotide Pyrophosphatase heterocyclic base, such asADP-ribose. Compounds with modifications of the adenine ring, as in the ethenoadenine analogs of NAD and AAD, were very poor substrates. Substitution of ethenoadenine for adenine would be expected to have very little effect on the chemistry of the pyrophosphate bond. Since the guanine analog of NAD was a suitable substrate for the enzyme, the inefficiency of the ethenoadenine derivatives is most likely due to steric factors rather than the loss of a hydrogen-bonding amino group. The bovine seminal fluid nucleotide pyrophosphatase catalyzed the hydrolysis of eNAD with kinetic constantsalmost identical to those obtained with NAD (20). The H. influenzue enzyme also displayed specificity in itsrequirement for a diesterified pyrophosphate group, since the phosphodiesterase substrate,bis-p-nitrophenylphosphate, and ADPdid not serve as substrates. A property displayed by the enzyme when acting on most substrates was non-Michaelian kinetics. Lineweaver-Burk plots were indicative of negative cooperativity. This property, first predicted by Koshland in 1968 (34), has been observed with several enzymes (35-40) including the sheep liver nucleotide pyrophosphatase (28). Hill coefficients ranged from 0.26 for NAD to values close to unity for several compounds which induced little or no cooperativity in the action of the enzyme. NAD was the strongest inducer of negative cooperativity. It has been proposed that theinteraction of the adenine ring of NAD with the NAD-binding site of sturgeon muscle glyceraldehyde-3-phosphate dehydrogenase is critical to the induction of negative cooperactivity in this enzyme (41, 42). A similar conclusion may be proposed for the H. infZuenzae nucleotide pyrophosphatase. All substrates functioned in a biphasic manner except e-NAD, eAAD, and nicotinamide guanine dinucleotide, compounds which have been modified in theadenine ring, relative to NAD. Substrates which lacked the pyridine ring were capable of inducing negative cooperativity, but the nature of the substituents on the ring, when present, also influenced the interaction of the substrate with the enzyme. AAD not only exhibited a much poorer binding affinity for the enzyme relative to NAD, but was almost devoid of the ability to induce cooperative interactions, as indicated by its Hill coefficient of0.93. Other analogs with various minor changes in the pyridine ring produced a wide range of Hill coefficients, indicative of widely different capabilites for induction of negative cooperativity. Glogger et al. (42) have proposed that the nicotinamide moiety of NAD is critical in orienting the adenine ring in the NAD-binding site of glyceraldehyde-3-phosphate dehydrogenase, and is therefore influential in the induction of negative cooperativity in this enzyme. , the enzyme activity AMP, when present at 20 p ~inhibited and abolished the negative cooperativity. On the basis of apparent dissociation constants, the enzyme showed a higher affinity for the adenine-half of NAD than for the nicotinamide-half. In addition, recognition by the enzyme of pyrophosphate moieties was manifested in the binding affinities for the adenine-containing compounds, i.e. ADP > AMP > adenosine. GMP was bound with a dissociation constant M not capable of abolishcomparable to AMP, but at20 ~ L was ing the biphasicity observed in double-reciprocal plots. This observation is in accord with the non-cooperative functioning of nicotinamide guanine dinucleotide as a substrate. The binding of adenine nucleotides was observed fluorimetrically and indicated that thebinding of these compounds occurred in at least two different modes. Dissociation constants from these data also reflected recognition of the pyrophosphate region of substrates, asthe order of affinity for the enzyme was ADP > AMP > adenosine.
6021
The nucleotide pyrophosphatase existed as a single polypeptide undernative and denaturing conditions. Because most, but notall, enzymes which display cooperativity consist of multiple subunits which interact to produce cooperative action (43), alternatemodels for the nucleotide pyrophosphatase were considered. The nucleotide pyrophosphatase from Phaseolus aureus exists as a dimer with a molecular weight of 65,000, but in the presence of AMP is converted to a tetramer (44).Gel-filtration high performance liquid chromatography studies indicated that in the presence of saturating concentrations of NAD, the H. influenzue enzyme may undergo a dimerization process. The negative cooperativity might reflect the transitionof this enzyme from a monomer to a dimer with concomitant conformational changes affecting enzyme function. A similar model has been proposed for the negative cooperativity of orotidylate decarboxylase (36). Alternatively, a large change in the Stokes radius could produce similar results. Further experimentation is needed to distinguish between these two possibilities. Conway and Koshland (45) have suggested that a benefit of negative cooperativity, opposite to the case of positive cooperativity, is that flux through negatively cooperative enzymes is relatively insensitive to large changes in substrate concentration.The constantturnover of extracellular NAD, leading to a constant internal supply of the dinucleotide, in the presence of the uncertain and fluctuating amounts ofNAD most likely encountered by this parasitic organism, would bebeneficial. Arginine and lysine residues are prime candidates for interactions with negatively charged pyrophosphate-containing compounds. The enzyme lost activity in a first-order process in the presence of 2,3-butanedione, a reagent that shows a preference for arginyl residues. ADP, at 20 p ~ only , partially protected the enzyme from inactivation. A time-dependentinactivation of the enzymewas also accomplished by exposure of the enzyme to Woodward’s Reagent K. ADP, at 20 p ~accelerated , this inactivation process, suggesting an ADP-induced conformational change making the essential carboxyl groups more susceptible to covalent modification. This observation is consistent with the observed recognition by the nucleotide pyrophosphatase of adenyl moieties for the induction of functional changes in theaction of the enzyme which are manifested as negative cooperativity. Growth studies indicated that H. influenzae grew well on enriched media supplemented with NAD at concentrations greater than 0.1 pg/ml. Equivalent concentrations of NMN were not aseffective as NAD in serving as V-factor. The rates of growth obtained with saturating concentrations of NMN were well below those obtained with saturating concentrations of several dinucleotides that served as V-factor. The lack of obvious saturation effects with NAD as V-factor is consistent with the negative cooperativity of the nucleotide pyrophosphatase, which allows for the functioning of the enzyme over a wide range of substrate concentrations. Growth with NMN as V-factor was not affected by AMP, whereas growth of the organism with NAD as V-factor was susceptible to inhibition by AMP. The functioning of €-NAD as V-factor was of particular interest. This analog would, subsequent to hydrolysis at the pyrophosphate bond, produce NMN in a manner similar to the use of nicotinamide hypoxanthine dinucleotide. Unlike the hypoxanthine analog, the ethenoadenine analog was very poor in serving as V-factor. It is possible that the decreased activity of €-NAD is due to the poor ability to serve as a substrate for the nucleotide pyrophosphatase. The hypoxanthine analog is a good substrate for this enzyme and also readily serves as V-factor. This situation is analogous to the
H. influenzae Nucleotide Pyrophosphatase
6022
use of AAD as a growth inhibitor. AAD was the most effective growth inhibitor analyzed, whereas E-AADwas much less effective, presumably due to its decreased efficiency in interacting with the nucleotide pyrophosphatase. The ability of H. influenzaeto use NMN asV-factor implies the existence of a pyridine nucleotide cycle that could lead to the synthesis of internal NAD. A functional pyridine nucleotide cycle in H. influenzae would not be expected to include steps involving nicotinamide or nicotinic acid, as these compounds are ineffective at functioning as V-factor. Recently, several microorganisms have been shown to utilize such a cycle, the PNC IV (46-50). It has been reported that when one of these organisms, Salmonella typhimurium, was genetically manipulated to rely totally on the PNC IV as a source of pyridine nucleotide, the organism was capable of using deamidated derivatives of NAD such asnicotinic acid adenine dinucleotide as asource of pyridine nucleotide (51). This is in contrast with observations with H. influenzae,which was not capable of converting deamidated derivatives of NAD to pyridine nucleotide. This observation, coupled with the ability of cell-free extracts of H. influenzae to catalyze synthesis of NAD directly from NMN as described in this report,implies that the organism satisfies its requirement for pyridine nucleotide by use of a PNC I1 (a two-membered pyridine nucleotide cycle). This cyclewouldinvolve the hydrolysis of external NAD to NMN followed by the uptake of NMN, or a dervative of it, with synthesis of NAD intracellularly from NMN. The ability to inhibit the growth of H. influenzae by specifically targeting the unique NAD metabolism of the organism has clinical implications. Problems such as the rise of antibiotic-resistant strains of the organism, increasing numbers of H. influenzae type b meningitis in adults (52), and reports of contagious H. influenzae infections (53) necessitate the development of alternate methods of therapeutic treatment. The present study documents the importance of the nucleotide pyrophosphatase of H. influenzae in processing extracellular NAD to meet the V-factor growth requirement of this organism. In this respect, selective inhibition or inactivation of this enzyme could serve as a focal point for controlling growth of the organism. Although AAD was the most effective growth inhibitor investigated, its use as a therapeutic agent would be problematic due to the neurotoxicity of aminopyridines (54). Although questions may arise regarding the rapid turnover of the adenine nucleotides to compounds ineffective at inhibition of growth, it is noted that these compounds are now in use for treatment of Herpes zoster infections (55). REFERENCES 1. Turk, D. C., and May, J. R. (1967) Haemophilus inflfluenzue: Its Clinical Importance, English Universities Press, London 2. Gunn, B.A., Woodall, J. B., Jones, J. F., and Thornsberry, C. (1974) Lancet 11,845 3. Syriopoulou, V., Scheifele, D., Smith, A.L., Perry, P. M., and Howil, V. (1978) J. Pediatr. 97,421-424 4. Lwoff, A,, and Lwoff, M. (1937) Proc. R. SOC.Lond. B Biol. Sci. 122,352-359 5. Schlenck, F., and Gingrich, W. (1942) J. Biol. Chem. 1 4 3 , 295296 6. Gingrich, W., and Schlenck, F. (1944) J. Bacteriol. 47, 535-550 7. Leder, I. G., and Handler, P. (1951) J. Biol. Chem. 189,889-899 8. Shifrine, M., and Biberstein, E. L. (1960) Nature 187, 623 9. Kahn, D. W., and Anderson, B. M. (1983) Biochemistry 2 2 , 8 a 10. Yost, D. A., Tanchoco, M. L., and Anderson, B. M. (1982) Arch. Biochem. Biophys. 217, 155-161 11. Barrio, J. R., Secrist, J. A., 111, and Leonard, N. J. (1972) Proc. Natl. Acad. Sci. U. S. A. 6 9 , 2039-2042 12. Malamy, M. H., and Horecker, B.L. (1964) Biochemistry 3 , 1889-1893
13. Specifications and Criteria for Biochemical Compounds, National Academy of Sciences (1979), National Research Council Publication 719 14. Rodden, J. L., and Scocca, J. J. (1972) Arch. Biochem. Biophys. 153,837-844 15. Bradford, M. M. (1976) Anal. Biochem. 7 2 , 248-254 16. Weber, K., and Osborn, M. (1969) J. Biol. Chem. 244.4406-4412 17. Witholt, B. (1971) Methods Enzymol. 18B, 813-816 18. Ainslie, G.R., Jr., Shill, J. P., and Neet, K. E. (1972) J. Biol. Chem. 247, 7088-7096 19. Foster, J. W., and Moat, A. G. (1980) Microbiol. Rev. 4 4 , 83-105 20. Buckon, M.E., and Anderson, B. M. (1980) Arch. Biochem. Biophys. 202,396-404 21. Haroz, R. K., Twu, J. S., and Bretthauer, R.K. (1972) J. Biol. Chem. 247,1452-1457 22. Swartz, M. N., Kaplan, N. O., and Lamborg, M. F. (1958) J. Biol. Chem. 232,1051-1063 23. Bischoff, E., Tran-thi, T., andDecker, K. (1975) Eur. J. Biochem. 51,353-361 24. Davies, R., and King, H. K. (1978) Biochem. J. 175, 669-674 25. Anderson, B. M., and Lang, C.A. (1966) Biochem. J. 101, 392396 26. Kumar, S. A., Rao, N. A., and Vaidyanathan, C. S. (1965) Arch. Biochem. Biophys. 11 1,646-652 27. Schliselfeld, L. H., Van Eys, J., and Touster, 0.(1965) J. Biol. Chem. 240,811-818 28. Krishnan, N. and Rao, N. A. (1972) Arch. Biochem. Biophys. 149,336-348 29. Nakajima, Y., Fukunga, N., Sasaki, S., and Usami, S. (1973) Biochim. Biophys. Acta 2 9 3 , 242-255 30. Glaser, L.,Melo, A., and Paul, R. (1967) J. Biol. Chem. 242, 1944-1954 31. Corder, C.N., and Lowry, 0.H. (1969) Biochim. Biophys. Acta 19 1,579-587 32. Ragab, M.H., Brightwell, R., and Tappel, A. L. (1968) Arch. Biochem. Biophys. 1 2 3 , 179-185 33. Clayton, R.A., and Hanselman, L. M. (1960) Arch. Biochem. Biophys. 8 7 , 161-166 34. Koshland, D. E., Jr., Nemethy, G., and Filmer, D. (1966) Biochemistry 5, 365-385 35. Apps, D. K. (1975) Eur. J. Biochem. 55,475-483 36. Brown, G. K., Fox, R. M., and O’Sullivan, W. J. (1975) J. Biol. Chem. 2 5 0 , 7352-7358 37. Meunier, J.-C., Buc, J., Navarro, A., and Ricard, J. (1974) Eur. J. Biochem. 49,209-223 38. Dalziel, K., and Engle, P.(1968) FEBS Lett. 1 , 349-352 39. Carvajal, N., Fernandez, M., Rodriguez, J. P., and Martinez, J. (1982) Biochim. Biophys. Acta 701,408-409 40. Bale, J. R., Chock, P. B., and Huang, _. C. Y. (1980) . . J. Biol. Chem. 255,8424-8430 41. Schlessinger, J., and Levitsky, A. (1974) J. Mol. Biol. 8 2 , 547561 42. Glogger, K. G., Balasubramanian, K., Beth, A. H., and Park, J. H. (1982) Biochim. Biophys. Acta 706,197-202 43. Neet, K. E. (1983) in Contemporary Enzyme Kinetics and Mechanism (Purich, D. L.,ed) p. 267, Academic Press, New York 44. Reddy, A. R. V., Ananthanaraynan, V. S., and Rao, N. A. (1979) Arch. Bwchem. Biophys. 198,89-96 45. Conway, A., and Koshland, D. E., Jr. (1968) Biochemistry 7, 4011-4023 46. Fyfe, S. A., and Friedman, H. C. (1969) J. Biol. Chem. 244,16591666 47. Imai, T. (1973) J. Biochem. (Tokyo) 7 3 , 139-153 48. Friedman, H. C. (1971) Methods Enzymol. 18B, 192-197 49. Kinney, D. M., Foster, J. W., and Moat, A. G. (1979) J. Bacteriol. 140,607-611 50. Manlapaz-Fernandez, P., and Olivera, B. M. (1973) J. Biol. Chem. 248,5150-5155 51. Foster, J. W., Kinney, D. M., and Moat, A. G. (1979) J. Bacteriol. 137,1165-1175 52. Spagnuolo, P. J., Ellner, J. J., Lerner, P. I., McHenry, M.C., Flatauer, F., Rosenberg, P., and Rosenthal, M. S. (1982) Medicine 6 1 , 74-85 53. Anonymous (1981) Lancet I, 649 54. Yeh, J. Z., Oxford, G. S., Wu, C. H., and Narahashi, T. (1976) J. Gen. Physiol. 68,519-535 55. Sklar, S. H., Blue, W. T., Alexander, E. J., and Bodian, C.A. (1985) J. Am. Med. Assoc. 253, 1427-1430
H. influenzae Nucleotide Pyrophosphatase Supplementary Naterial To:
6023
G e l filtration HPLCwas done usrnga Bio-Rad TSK-250column (7.5 x 300 M sodium Sulfateon a Spectra-Physics SP35OOB liquid chromatograph equipped with a Spectra-Physics model 770 detector set at 233 nm. The column was maintained 33-c at with a flow rate of1.0 ml/min.
m ) equilibrated in0.1
CHARACTERIZATION
OF
HAENOPHILUS
INFLUENZAE
NUCLEOTIDE
PYROPHOSPHATASE
by Amino acid analyses were performed using a Beckman HPLC amino analyzer equipped with a sodium cation exchange column obtained Pickering.
David W. Kahn and Bruce N. Anderson
acid from
RESULTS
EXPERIMENTAL PROCEDURES purification
Naterials
of
the
Nucleotide
PW-ODhOSDhhataSe
E. Purification was initiated by preparing spheroplasts of frozen Haemopbilus influenzae strain Rd was obtained from Dr. W. L. Albritton I of broth culture using the procedure described influenzae cells from 4.5 of the University of Saskatchewan. Saskatoon. Brain Heart Infusion was in ExDerimental Procedures. Particulate matter was collected bu centrifuaaobtained from Fisher Scientific. Reagent grade Tris (hydroxymethy1)amino- ;ioi~~>&>-lO-~mii it~i9]OOo-x,at 4-c; The Supernatant (2.2 1 ) &as then methane, hemin, histidine, streptomycin sulfate, Sepbacryl S-200, lysozyme, dialyzed overnight against 2 x 4 1 of 50 m Tris-HCI, pH 8.5. The dialysate Woodward's Reagent K. bis-(g-nitrophenyl) phosphate, Coomassie Brilliant was then adjusted to 45% saturation of amonium sulfate at 4'C. The superBlue R. N,N,N',N'-tetramethylethylenediamine (TENED) and protein standards natant after centrifugation was adjusted to 65% saturation of ammonium were purchasedfrom sigma. All nucleosides, mono-and dinucleotides were was resuspended in60 ml of 50 mY Trir-HCI, sulfate. The resulting pellet purchased from Sigma except 3-aminopyridine adenine dinucleotide (AAD)'. pH 8.5 and dialyzed overnight against 1 2of 50 mp1 Tris-HCI. pH 8 . 5 at 4-C. nicotinamide I,N6-ethenoadenine dinucleotide (E-NAD), 3-aminopyridine The dialysatewas applied toa column (1.5 x 30 cml of phosphocellulose I. N6-ethenoadenine dinucleotide ( c - m J , and 3-methylpyridine adenine of equilibraequilibrated in 50mEl Tris-HCZ. pH 8.5. Two column volumes the (10,Il). Nicotin- tion buffer were used to remove unbound protein. One column volume of 0.2 dinucleotide. which were prepared by publisbed procedures amide, nicotinic acid.N,N'-methylenebisacrylamide and Coomassie Brilliant M sodium fluoride in equilibration buffer was washed through the column in Blue 0-250 were from Eastman Organic Chemicals. Natrex Green Gel A and an effort to elute alkaline DbOSDhataSe activitu from the column. The Natrex BlueGel A werefrom Amicon. Acrylamide was from Bio-Rad. 2,3-Butane- nucleotide pyrophosphatase w>s eiuted using a llnear gradient from 0.2-1.0 dione was from the Aldrich chemical Company. N KC1 in the equilibration buffer. Fractions (2.5 0 1 ) containing activity (Fig. I ) , assayed fluorimetrically, were pooled and dialyzed overnight against 2I of 50m Tris-HC1. pH 8 . 5 . C e l l s were grown in 750 ml of medium in 2.8 I Fernbach flasksat 37-C in a New Brunswick G-25R incubator with shaking at 120 cycles/min. The medium, 28 g of solid Brain Heart Infusion in 750 ml of distilledWater was autoclaved for15 min at 120°C under 15p.s.i. After cooling to room temperature. 7.5 ml of aNAD solution(300 p g / m l ) and 7.5ml of a hemin6 histidine solutionwere added. The hemin-histidine solution contained 10 2.0 50 mg each of hemin and histidine in 4.6 ml of distilled water 0.4 to mlwhich of triethanolamine was added and the mixture was placed a in Water bath at E 55*C for 10 min. After incubation, 5.0 ml of distilled water were added. II 40 c Both the NAD and the hemin-histidine solutions were added while undergoing 1.6 filter sterilization. Growthwas initiated by addition of an innoculum (3.0 ml) of late linear phase c e l l s adjusted to16% glycerol and stored at 0 1.2 -7OOC. m
'
1
->
.-
(v
2In studies of the nucleotide pyrophosphatase, cells were harvested by Centrifugation at 18.000 xa for I o min. washed twice in m 50 Tris-HCI, DH 8.5, resuspended in a minimil amount of theSame buffer and stored at . -15OC. Cells (2.0 gwet weight) from500 ml of broth were suspended in 5.0 ml of 50 m Tris-HCI. DH 9.0 for studies of extracellular NAD hudrolusis. One ml Of 5 m KgcI 'aid solid NAD(2 mH finai conceitratioil&e a&& At timed intervals,1.5 ml aliquots and the mixture incgbated at 37OC. were removedand centrifuged and the Supernatant solution was filtered using a0.2 micron filter. A sample of filtrate diluted With 200 m 20 40 60 80 I00 120 140 160 180 DOiaSSiUm DhOsDhate,OH 4.0. was analuzed bv ion-exchanae HPLC. For studies if NAD py~oph&phor;l>se, cells from i . 5 I of broth, "&e disrupted by FRACTION NUMBER at 100,000 sonication anda soluble fractionwas obtained by centrifugation x g for one hour at 4°C. Fig. 1. Phosphocellulose ion-exchange chromatography. At the first arrow, the colunvl was eluted with equilibration buffer containingmH 200 NaF. At In studies evaluating the ability of COmDOUndS to asServe V-factor. n KCI in equilibration the secondarrow, a linear gradient from 0.2-1.0 growth was initiated by addition0;s ofml Of-freshly grown cells(late buffer was applied. Enzyme activity w a s determined fluorimetrically. linear phase). The innocula (1%. v/vl in all experiments, except those where NHN was used a s v-factor. contained1 ,u _, o h l NAD. I n euoeriments with , NNN as V-factor, theimocuZa were grown in the presenceI pg/mI of NNN. 0.1 p g / m l of NAD and In growth inhibitionexperiments, the media contained of inhibitor in a volume50 of ml. the indicated amount The dialysate from above was applied toa column (1.2 8.0 x cm) of Lysozyme digestions were performed hy modification of the procedure Katrex.of Green Gel A that was equilibrated 50inm Tris-HCI, pH 8.5. Two Malamy and Horecker (12). The cell suspension was adjusted to 7 EDTA used to remove unbound orotein. column volumes of eouilibration buffer were and lysozyme wasadded to a final concentration of 3 p g / m l . This solution The enzyme vas elutld using a line&&dient~of 0-50O-&~KC7~In-~tbeepuiliwas incubated at37'C for one hour and then centrifuged at19.000 x g for bration buffer. Fractions (2.5 ml) containing activity (Fig.2) were I5 min. pooled and dialyzed overnight inI of 2 50 m Tris-HC1, pH 8.0. A fluorimetric assay was used to monitor nucleotide pyrophosphatase activity during purification. Reaction mixtures contained mH25Tris-HCZ, pH 8.5, 1 mH NgCla, ,300 nm FAD and enzyme aintotal volume of 3.0 ml. Fluorescence inte sity was determined at an excitation Wavelength of nm 465 and emission maximum of 510 nm using a Perkin-Elmer 650-40 fluorescence spectrophotometer. ~
~~
~
~
~~~
~~~
~
~
~
~
~~
~~~
~~~
~~~~
.~~~~~~~
was used for characterization of the purified A titrimetric assay enzyme. All pH-stat measurementswere made usinga Radiometer PHn82pH meter equipped with a GK232OC combination electrode, TTT80 titrator, and ABUIO autoburette. The end point fora l l titrations, except those involved on activity, was 8.0. Reaction in the investigation of the effect of pH mixtures contained50 mn KCI, substrate and enzyme a in total volume of 3.0
ml
Ig
.
-
NAD pyrophosphorylase activity was determined spectrophotometrically at 340 m using a Beckman ActaNVI spectrophotometer. Incubation mixtures contained 100 mEl Tris-HC1, pH 7.5, 15 m NgClz, 5 mEl ATP. 2.5 mn NNN and 0.2 ml enzyme in a total volumeof 1.0 ml at 37Qc. At timed intervals aliquots were removed, deproteinized with trichloracetic acid a and 0.2 m1 sample of the supernatant was then addedto a cuvette containing 0.88 m l 90 mN unbuffered Trisand 0.5 N ethanol. The absorbance change at 340 nm after addition of 7 units of yeast alcoho~dehydrogenase was used to c a ~ c u late the amountof NAD formed (13).
Ion-exchange HPLCwas performed usinga column (4.6 250 x mml packed with Alltech RSIL-AN resin, 5 micron particlesize, equilibrated in100 m potassium phosphate. pH 4.0. The flaw ratew a s 1.0 ml/min and the column was maintained at room temperature. Reverse-phase, ion-pair HPLCwas conducted usinga column (4.6x 250 m ) packed with Alltech RSIL-C18-HL resin witba particle size of5 microns. The columnwas equilibrated with 35 &?potassium pbosphate. 2.8mM tetrapropylammonium hydroxide and 30% metbanol adjusted topH 5.5. The column was maintained at 50-C with a flow a Spectra-Physics rate of 0.75 ml/min. Both HPLC analyses were done using SPBOOO chromatograph equipped with a Spectra-Physics model 770 variable wavelength detector. Absorbance of the column eluent w a s monitored at 260 m.
02
01
The 2',3'-cyclic phosphodiesterase was assayed according to published (15). procedures (14). Protein was determined by the method of Bradford SDS polyacrylamide gel electrophoresis was performed according to and Weber Osborn (16). 10 mg of NNN with 20 mg of Nir was prepared from NNN by incubating wheat germacid phosphatase (0.36u/mgl in 10 ml of 5m sodium acetate,pH oroduct was ourified bU 5.0 for 30 hours with constant stirrlno. The ion-exchange chromatography and w& id;ntifi=d' a; nicot;na;ide~;Ibo;;de by thin-layer chromatography (17).
O3
FRACTION NUMBER
Fig. 2. natrex Green el A affinity chromatography. The column was equilibrated in 50 mN Tris-HCI. pH 8.5. At the arrow, a linear gradient from 0-500 mN KC1 in the equilibration buffer w a s applied. Tbe dialyzed fractions from the Natrex Green Gel A step were applied to a column (1.2 x 8.0cm) of Natrex BlueGel A equilibrated in m 50 Tris-HCI, pH 8.0. Two column volumes of the equilibration buffer were applied to remove unbound protein. The enzyme w a s eluted f r o m the coIumn using a linear gradient of0-1 N KC1 in the equilibration buffer. Fractions containing nucleotide pyrophasphatase activity (Fig. 3) were pooled and dialyzed twice against 4 I of 200 mn KCI. In preparation for titrimetric was adjusted topH 8.0 by the analyses. The pH of the dialysis solution gradual addition of a dilute NaOH solution.
Pyrophosphatase Nucleotide H. influenzae
6024
r
I
I
I
I
I
I
The amino acid composition of the purified enzyme (Table I ) was determined witha Beckman HPLC amino acid analyzer equipped awith sodium cation exchange column obtained from Pickering Co.
I
90 TABLE Amino
70 enzyme moles/moleAcid
Acid
I Analysis
Amino
50 Lysine Arginine Histidine Aspartate Glutamate Serine Threonine Proline Cysteic acid Methionine Glycine AZanine Valine Leucine Isoleucine Tyrosine Phenylalanine
30
IO
IO
20
30
40
50
60
70
FRACTION NUMBER Fig. 3. Matrex ~ l u eGel A affinity chromatography. The column was eguilibrated in 50 rn Tris-HCI. pH 8.0. At the arrow,a linear gradient from0-1 M KC1 in equilibration buffer was applied.
Additional P r o ~ r t i e sof
the
Purified
Nucleotide
PyrophosDhatase
The productsof the nucleotide pyrophosphatase-catalyzed hydrolysis FAD wereidentified by reverse-phase, ion-pair HPLC. As shown in Pig.4 , the soleproducts observed were flavin mononucleotide ( F N N J and ANP.
44 13
14 72 58 40
34 24 1 IO 49 50 41 47
30 20
22 569
The thermostabilityof the purified enzyme was determined over the range from 15-5OoC. AS shown in Fig. 5 , activity was lost at each temperatUre a s a first order rate process. Half-lives of thermal denaturation at of these temperatures ranged from 110-19 min.
1
90 0 Ilme
FAD
80
-
70
60
10 minutes
5c
25 rninules
4c
3c
47 mlnutes
IO
20
30
40
TIME (rnin) Fig. 5 . Thermal denaturationof the nucleotide pyrophosphatase. Incubation mixtures containedI pg of the nucleotide pyrophosphatase and 50 mfl Tris-HCI. DH.. 8.0 At timed intervals,aliwots were = ~.. in a total volumeof 1.0 ml. rewved and assayedfor activity fluorimetrically. The tempirature of the incubation mixtures were: line 1. 15OC; line 2, 35'C; line 3 , 40'C; line
FMN
~
Fig. 4. Product analysis of the nucleotide pyrophosphatase-catalyzed hydrolysis of FAD Dy reverse-phase ion-pair HPLC. Incubation mixtures contained 2s mfl Tris-HCl PH 8 . 5 mfl Ngcl 333 ~ r and n 2 pg of the nucleotide pyr0phosphata;e ina iota1 vozu9 of 3 . 0 ml at 3 7 0 ~ .
4, 50°C.
i
The effect of temperature on the rate of the enzyme-catalyzed reaction investigated over the range from 5.4-45.8-C. These data are presented in Fig. 6 as an Arrhenius plot. The data were linear over the entire 8.2 kca>/mol for the temperature range observed and an activation energy of enzyme-catalyzed reaction was calculated.
was
Nucleotide H. influenzae
Pyrophosphatase
6025
w a s inactivated by 2,3-butaneThe purified nucleotide pyrophosphatase dione. At 100 pH 2.3-butanedione, the enzyme lost activity with a half-life Addition of 20 pM ADP to an identical incubation mixture orovided minimal arotection from this arocess. The enzume Was also inactivated by WoodwardAr; Reagent K, a reagent which modifies-reactive carboxyl of 50 M Woodward's groups. As shown inFig. 8, inactivation in the presence Reagent K proceeded witha half-life of 36.0 min. Addition of 20 pH ADP to an identical incubation mixture caused an acceleration of the loss of activity, producinga half-life of 14. 7 min. of 21.8 min.
.
I oc
90 8C 70
6C t
> -
5c 4c
a
L S
3c
20
I
1
I
310
320
330
340
1
I
350
360
I
I
I
I
I
IO
20
30
40
50
TIME (min)
f (OK x1051
Fig. 8 . Inactivation of the nucleotide pyrophosphatase with Woodward's Reagent K (WR-K). Incubation mixtures contahed~25mn Tris-HCI. pH 8.5, the indicated amount of WR-K1 pand g nucleotide pyrophosphatase in a total volume of 1.0 ml at 15'C. At timed intervals. aIiouots were removed and ~ i g .6. The effectof temperature on the nucleotide pyrophosphatasehydrolysis of NALL dctivity was determined using the titrimetricassayed for activity using the fluorimetric aisay.. The reactions contained: line I , 2 mM WR-R; line2, 10 mM WR-K; line 3 , 50 M FIR-& line 4, 50 mM assay at a NAD concentrationof 188 !JM. WR-K + 20 pM ADP.
The intrinsic fluorescence of the purified nucleotide pyrophosphatase was quenched by interactions with adenine nucleotides. Data obtained on the bindingof AMP is shown inFlg. 7.
300
240
I 20
60
40
80
I
I
I
120
160
200
Fig. 7. Quenching of the intrinsic fluorescence of the nucleotide pyrophosnhatase with M P . Cuvettes containino25 mn Tris-XC1, pH 8.0 and9.0 p g of
