Hydrolysis of Urea by Ureaplasma urealyticum Generates a
Transmembrane Potential with Resultant ATP SynthesisD. G. E. SMITH,* W. C. RUSSELL, W. J. INGLEDEW, AND D. THIRKELL
Division of Cell and Molecular Biology, School ofBiological and Medical Sciences, University of St.Andrews, Irvine Building, North Street, St. Andrews, Fife, KY16 9AL Scotland, United Kingdom
Received 30 December 1992/Accepted 17 March 1993
When urea is added to Ureaplasma urealyticum, it is hydrolysed internally by a cytosolic urease. Under our
measuring conditions, and at an external pH of 6.0, urea hydrolysis caused an ammonia chemical potentialequivalent to almost 80 mV and, simultaneously, an increase in proton electrochemical potential (h) of about24 mV with resultant de novo ATP synthesis. Inhibition of the urease with the potent inhibitor flurofamideabolished both the chemical potential and the increase ofAp such that ATP synthesis was reduced to -5% ofnormally obtained levels. Uncouplers of electrochemical gradients had little or no effect on these systems. Theelectrochemical parameters and ATP synthesis were measured similarly at three other external pH values. Anychange in Ap was primarily via membrane potential (A*), and the level of de novo ATP synthesis was relatedto the increase in Ap generated upon addition of urea and more closely to the ammonia chemical potential.Although the organisms lack an effective mechanism for internal pH homeostasis, they maintained a constantApH. The data reported are consistent with, and give evidence for, the direct involvement of a chemiosmoticmechanism in the generation of around 95% of the ATP by this organism. Furthermore, the data suggest thatthe ATP-generating system is coupled to urea hydrolysis by the cytosolic urease via an ammonia chemicalpotential.
Ureaplasma urealyticum is a small, wall-less, free-livingprokaryote which has been associated with infections of thehuman urogenital tract (2) and, more recently, with diseaseof preterm low-birth-weight neonates (4). The organismpossesses enough genetic material for around 500 geneproducts (11), and it requires a very complex medium tosupport growth. Since it has no cytochromes and apparentlylacks quinones (17), oxidative phosphorylation does notoccur or is severely limited; it also lacks enzymes to utilizea number of substrates (15, 16, 27) and has an incompletespectrum of tricarboxylic acid cycle enzymes (3). Thus,many mechanisms of substrate-level phosphorylation canalso be ruled out.Although one investigation has shown that an FoF1 pro-
ton-translocating ATPase is conserved in mycoplasmas (30),U. urealyticum was not included in that study. Others haveshown, however, that the organism has membrane-boundATPase with similarities to the FoF1 ATPases of otherbacteria (22), and it has been reported that ATP determina-tion represents a reliable and accurate means of measuringgrowth (28). It is therefore attractive to propose that an FoF1membrane ATPase in U. urealyticum plays an important rolein energy generation. Such a proposal could rely upon achemiosmotic mechanism (10), as suggested elsewhere (9),whereby generation of a transmembrane electrochemicalpotential would provide the electromotive force for protonsto enter the cell via the FoF1 ATPase to generate ATP. Suchelectrochemical potentials may also be utilized in transportprocesses. It has been suggested that since U. urealyticumhas a potent cytosolic urease, urea hydrolysis to ammoniumions and carbon dioxide generates a transmembrane poten-tial which drives ATP synthesis (9). While it has beenreported that urea and urease activities are essential for ATP
* Corresponding author.
synthesis (19-21), the precise nature of any interrelationshipremained uncertain.
In this study, we have examined the influence of ureahydrolysis upon electrochemical parameters and the influ-ence of these parameters upon ATP synthesis.
MATERIALS AND METHODSUreaplasma strain. U. urealyticum, serotype 8 (T960), was
a gift of D. Taylor-Robinson (Clinical Research Centre,Harrow, England).
Chemicals. Radiolabelled chemicals were obtained fromDu Pont, U.K., Ltd., Soluscint 0 and Solusol were fromNational Diagnostics, flurofamide was a gift of I. Kahane(Hebrew University, Hadassah Medical School, Jerusalem,Israel), and, except where stated, all other chemicals werefrom Sigma.
Cell culture and harvest. U. urealyticum was cultured(500-ml cultures) and harvested as described previously (18).After cell harvest, the pellets obtained by centrifugation(25,000 x g, 20 min) were washed with 0.25 M NaCl, and thefinal pellets were resuspended in an appropriate buffer (seebelow) at -1 mg of protein ml-' (estimated by the method ofLowry et al. [8]). On the basis of the determination ofcolor-changing units per milliliter (18), no apparent alterationin viability of the cells was apparent after this treatment.
Determination of intracellular volume. After washing, thepellet was suspended in 150 mM choline chloride-80 mMNa2SO4-50mMMES [2-(N-morpholino) ethanesulfonic acid]buffer (pH 6.0) containing tritiated water (1.0 ,Ci ml-';specific activity of stock, 2.5 mCi ml-') or tritiated inulin(1.0 ,Ci ml- ; specific activity of stock, 100 mCi g-1).Aliquots of suspension (200 ,u) were layered onto 100 ,u ofsilicone oil (DC 550 [BDH]; bis-3,5,5-trimethyl hexyl phtha-late [Fluka], 60:40 [vol/vol]) in Eppendorf tubes (in tripli-cate), incubated (37°C, 15 min), and then centrifuged (12,000x g, 5 min). Aliquots (100 RI) of aqueous phase were
removed and added to scintillant (Soluscint O-Triton X-100,2:1 [vol/vol]), and the remaining pellets were solubilized inSolusol and placed in a scintillation vial to which scintillantwas then added (Soluscint O-Triton X-100, 9:1 [vol/vol]).The internal water volume was calculated as describedpreviously (14).Measurement of membrane potential (AJ) and transmem-
brane pH difference (ApH). Washed cell pellets were sus-pended in buffer (150 mM choline chloride, 80 mM Na2SO4,and either 50 mM HEPES [N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid] or 50 mM MES). Buffer with MESwas adjusted to pH 5.5, 6.0, or 6.5, and buffer with HEPESwas adjusted to pH 7.0 with 1 M NaOH. Aliquots of thesuspension (100 Al) were added to Eppendorf tubes contain-ing 100 11l of silicone oil overlaid with 100 p.l of buffercontaining radiolabelled probe and, where required, urea togive a final concentration of 10 mM. Ad was monitored byusinf tritiated tetraphenyl phosphonium bromide (0.1 ILCiml-; concentration in assay, 2.2 x 10-12 mol ml-'), andApH was monitored by using a variety of probes (tritiatedacetic acid, 0.4 p.Ci ml-' [concentration in assay, 1 x 10-'mol ml-1]; [14C]chloroacetic acid, 0.2 tiCi ml-' [concentra-tion in assay, 0.6 x 10-7 mol ml-l]; [14C]methylamine, 1.0p.Ci ml-' [concentration in assay, 2.6 x 10-7 mol ml-1]; or[14C]ethanolamine, 1.0 p.Ci ml-' [concentration in assay, 2.1X 10-7 mol ml-l]). Suspensions with probes were incubated(370C, 30 s) and then centrifuged as described above. Theaqueous phase and cell pellet were treated and radioactivitywas determined as described before. Corrections for thenonspecific binding of probes to cellular components werecarried out by inclusion of controls with suspensions of cellslysed by sonication (three times, 10 s each time, on ice; MSESoniprep 150) at the same protein concentrations (mg ml-').These controls contained no viable cells upon culture, andwe had demonstrated previously by electron microscopy(data not shown) that sonicated cells were disrupted. Theytypically bound -10% of the total radioactivity of viablecells in the assays. Each time, assays were carried out intriplicate. Calculations of Ali and ApH were described else-where (23).Measurement of internal and external ammonia (NH3 and
NH4+) concentrations. The basis of the method for measuringinternal and external ammonia concentrations was essen-tially the same as that used for determination of Adi and ApH,and again, on each occasion, the assay was carried out intriplicate. To an Eppendorf tube containing 100 Atl of siliconeoil layered over 50 Atl of trichloroacetic acid (TCA; 15%[wt/vol]), first 100 A.l of ureaplasma suspension in buffer wasadded, and then 100 plI of buffer containing 20 mM urea wasadded. After incubation (370C, 30 s), the tubes were centri-fuged as described above. Under these conditions, ureaplas-mas were pelleted through the oil into the TCA and lysed,enzymes were inactivated, and intracellular ammonia wasreleased. Aliquots (10 p.l) of the upper aqueous phase wereremoved and added to 90 A.l of solution A (0.5 M NaOH,3.3% [wt/vol] TCA) for measurement of external ammonia.After the remainder of the aqueous phase and the oil werecarefully aspirated, aliquots (20 ALl) of the lower TCA phasewere removed and added to 80 RI of 0.56 M NaOH formeasurement of internal ammonia. Thus, all samples con-tained a final concentration of 0.45 M NaOH and 3% (wt/vol)TCA. This procedure was necessary because we found thatthe Bertholet assay for determination of ammonia was pHsensitive (data not shown). The Bertholet assays were car-ried out in a microtiter plate, and, to each well, 50 p.l ofphenol nitroprusside and 50 pl of alkaline hypochlorite were
added (Sigma urea nitrogen colorimetric kit for the Bertholetreaction). After incubation (20'C, 20 min), A6o0s were deter-mined (Titertek Multiscan). Ammonia concentrations weredetermined from a standard curve prepared similarly fromappropriate concentrations of NH4Cl.
Determination of intracellular ATP. Measurements of in-tracellular ATP were carried out in duplicate by usingureaplasma cell pellets suspended in MES or HEPES bufferas described above but containing 2 mM MgCl2 and 10 mMNaH2PO4. Assays were performed in the presence andabsence of exogenous 40 mM urea at external pH values of5.5, 6.0, 6.5, and 7.0. In addition, assays were performed atan external pH of 6.0 in the presence and absence ofexogenous 40 mM urea and in the presence of a range ofconcentrations of uncouplers which included those usuallyemployed against other bacteria (200 p.M 2,4-dinitrophenol[DNP]; 200 p.M gramicidin; 10.0, 40.0, 160.0, and 640.0 p.Mcarbonyl cyanide m-chlorophenylhydrazone [CCCP]; or ure-ase inhibitor [flurofamide, 25 p.M] or the FoF1 ATPaseinhibitor NN'-dicyclohexylcarbodiimide [DCCD, 0.01 and0.001 mM]). Appropriate ureaplasma suspensions were in-cubated (370C, 30 s [except where stated]) and then adjustedto 10% (vol/vol) with respect to TCA. After dilution (1:50[vol/vol] in 100 mM Tris-acetate [pH 7.5] containing 2 mMEDTA), ATP was measured by using a luciferin-luciferasekit (Bio-Orbit) on a model 1250 luminometer (Bio-Orbit).ATP was calculated by using standard ATP solutions andexpressed as nanomoles of ATP per milligram of cell protein.
RESULTS
Determination of intracellular water volume. Under ourassay conditions, the internal volume of U. urealyticum was3.8 ± 0.2 ,ul mg of cell protein-'. This is comparable toreported volumes of 1.6, 2.5, and 4.8 p.l mg of cell protein-for Mycoplasma gallisepticum, Acholeplasma laidlawii, andMycoplasma mycoides var. capri, respectively (6, 23, 24).This value was then used in the calculation of internalconcentrations of probes and of ammonia.A* and ApH determinations. Throughout the assays, all
ureaplasma cells appeared to pass through the silicone oilmixture. There was no visual evidence of residual material inthe aqueous phase, and no evidence of viable cells could befound in the aqueous phase by culture. At the density of theoil mixture used, even membranes would be expected topass through the oil. Neither methylamine nor ethanolamine(accumulated by cells with a more acidic internal pH thanthe external environment) was accumulated by ureaplasmasat any of the four external pH values investigated. Ethano-lamine was also used as a control in case methylamine couldbe transported by an ammonia carrier. This indicated thatthe internal cellular pH was higher than that of the externalbuffers used. Confirmation of this was obtained by uptake ofacetic acid and chloroacetic acid at each external pH, givingreproducible ApH values over a large number of repeatassays, each carried out in triplicate. Results (Table 1) showthat both A* and ApH values not only varied with externalpH but also were elevated in cells catabolizing urea. Theseeffects are discussed later. Under these circumstances, thevalues of A* rose between an external pH of 5.5 to 6.5 andthen fell at pH 7.0. The differential in AqJ values with andwithout added urea, however, was greatest at external pH6.0, showing decreased values below and above this pH.Conversely, the values for ApH in these cells displayed anarrower range and were raised only minimally in thepresence of urea. Assays carried out with 100 mM phosphate
TABLE 1. Determinations of ureaplasma A* and ApH in the presence and absence of exogenous 40 mM urea and in the presence of40 mM urea plus 25 ,uM flurofamide at four external pH values
a Values are means + standard errors of the means.b A, in the absence of urea; B, in the presence of exogenous 40 mM urea; C, in the presence of 40 mM urea plus 25 p.M flurofamide.
buffer, as applied elsewhere (19, 21), gave similar valueswithin the standard error. Inclusion of uncouplers (CCCP,DNP, gramicidin) over a range of concentrations, or ofuncoupler solvent (ethanol), generally failed to alter thevalues obtained for either A* or for ApH with or withoutadded urea (Table 2). This was peculiar to the ureaplasmassince the ionophores were active in parallel assays with astrain of Escherichia coli (data not shown). A significantreduction in Ai values was seen, however, with higherconcentrations (160.0 and 640.0 raM) of the proton ionophoreCCCP, although at these concentrations its effect may benonspecific. On the other hand, inclusion in the assays of thepotent urease inhibitor flurofamide (5) at 25 p.M totallyabolished any increase in either A* or ApH observed in thepresence of exogenous urea (Tables 1 and 2). This showedthat at least the increased A* and ApH values observed inthe presence of urea, compared with those in its absence,were directly related to urease activity.
Internal pH of U. urealyticum. From the values of ApH, the
TABLE 2. A* and de novo ATP synthesis determined for U.urealyticum, at an external pH of 6.0, in the presence and
absence of uncouplers, or the urease inhibitor flurofamide, orthe F0F1 ATPase inhibitor DCCD0
ATP synthesizedUncoupler or inhibitor A/$ (mV)b (nmol mg of cell
0.01 mM ND 3.34 ± 0.400.001 mM ND 4.72 ± 0.61a De novo ATP synthesis was measured in the presence of 40 mM urea
minus the value determined in the absence of added urea. The assay periodwas 30 s. In the presence of DCCD, cells were incubated for 3 min at 20'Cprior to the addition of urea for the assay of ATP.
b Values are means + standard errors of the means.C Values are means, with ranges indicated.d ND, not done.
intracellular pH can be calculated. This was shown to varydirectly with extracellular pH and found to be 0.78 to 0.95pH unit higher than the pH of the external buffer. Thisfinding was reproducible in many assays and suggests that,like other mycoplasmas (1, 7), U. urealyticum lacks aneffective mechanism of internal pH homeostasis but main-tains a relatively constant ApH.Transmembrane chemical potential of ammonia. The con-
centrations of ammonia internally and externally were mea-sured during urea hydrolysis under the same conditions (30-sincubation) as those used for the measurement of A4 andApH (see Materials and Methods). Although the internalammonia concentrations varied with external pH under theconditions applied, the external ammonia concentration was3.1 + 0.1 mM irrespective of external pH or of internalammonia concentration. A concentration difference of am-monia (inside to outside the cells) of -21-fold, equivalent to78.9 mV (calculated as described previously [14]), was seenat an external pH of 6.0. At external pH values of 5.5, 6.5,and 7.0, the values were equivalent to 69.9, 74.2, and 58.6mV, respectively. These differences were abolished if theurease inhibitor flurofamide was added at a concentration of25 ,uM in the assays.
Intracellular ATP synthesis. A time course study of ATPsynthesis was carried out at an external pH of 6.0. Aminimum assay period of 30 s was used to correlate thesedata with electrochemical data (see Discussion). The results(Table 3) indicated that a 30-s incubation gave maximalvalues over the time course investigated, that any endoge-nous urea in the cells was rapidly depleted, and that with a30-s incubation, ATP synthesis from endogenous urea hy-drolysis and/or substrate-level phosphorylation made a con-tribution of only -5% to the values determined in thepresence of exogenous urea. The decline in ATP valuesdetermined with time of incubation in both the presence and
TABLE 3. Time course of ATP production by U. urealyticum atan external pH of 6.0 in the presence and absence of exogenous
40 mM ureaa
Assay incubation ATP produced (nmol mg of cell protein-l)btime (s) +Urea -Urea
a De novo ATP synthesis is the value determined in the presence of 40 mMurea minus the resting ATP level measured in the absence of urea.
b Values are means, with ranges indicated.
absence of exogenous urea presumably is a consequence ofthe rate of ATP utilization being greater than the rate ofsynthesis as the available urea decreases through hydrolysisand as the ammonia chemical potential dissipates.De novo ATP synthesis occurred at all four external pH
values in the presence of exogenous urea (Table 4). Theamount ofATP measured is consistent with the electrochem-ical data, in particular, with the magnitude of the differentialswith respect to Ap in the presence and absence of urea(Table 5) and the ammonia chemical potential.The effect of uncouplers on ATP synthesis at an external
pH of 6.0 (Table 2) paralleled the effect of the same uncou-plers on measured Ad (and hence Ap) values. Synthesis ofATP was, however, reduced to -5% of its maximum valuewhen 25 jxM flurofamide was included in the assays. TheFoF1 ATPase inhibitor DCCD also significantly inhibitedATP synthesis (Table 2).
DISCUSSIONIt has been suggested that the concomitant activity of
ureaplasma urease and an ATPase must occur to permit ATPsynthesis (21). Others have hypothesized that urea hydroly-sis produces an electrochemical gradient to generate ATP bya chemiosmotic mechanism (9, 21). In these experiments, wehave attempted to unravel the mechanism(s) which may playa role in such a system.
All electrochemical parameters were determined by usingiso-osmotic buffers and careful sedimentation and suspen-sion because of the known osmotic fragility of these cells.With these precautions, variations in both Al and ApHvalues were minimized. Moreover, for the measurement ofA* and ApH, it is necessary to maintain a stable external pH.Although ammonia is released extracellularly by ureaplas-mas after hydrolysis of urea by cytosolic urease, no increasein external pH was measurable during the 30-s period of the
TABLE 5. Determination of ureaplasma proton electrochemicalpotential (Ap) in the presence and absence of exogenous 40 mM
urea and in the presence of 40 mM urea plus 25 p.M flurofamide atfour external pH valuesa
AD (mVl)External DifferentialpH No urea +40 mM urea +25 A±M Ap(B-A)
assays. The parameters were all maximal at -30 s after theaddition of urea, the minimum assay period possible with ourprotocol. This reflects the pulsed nature of the experimentalsystem and may correlate with the in vivo situation wherenot only would cells receive an intermittent nutrient supplybut also released metabolites would be quickly dissipated.The data showed that the addition of urea to viable
ureaplasmas produced an increase in both At and ApH. Thereported increase in Ap may be too low to account for the -3mM concentration increase in ATP determined. However,since the relationship between Ap and ATP synthesis isnonlinear, this rise may increase the energization of ure-aplasmas sufficiently to initiate ATP synthesis dependentupon atypical H+- or monovalent cation+-to-ATP stoichiom-etry as observed in other microorganisms. In addition, theAp values determined in the absence of urea suggest gener-ation by urea-independent "housekeeping" metabolic pro-cesses. Variation in these Ap values with changing externalpH suggests that these processes are pH dependent and mayreflect an energy requirement for maintenance of cellularviability under suboptimal conditions. Although very diffi-cult to prove experimentally, we suggest that urea hydrolysisdown-regulates some or all housekeeping processes (perhapsin a pH-dependent manner) either directly by urea or via theammonia chemical potential. If this was the case, the mag-nitude of Ap determined in the presence of exogenous ureaand not the differential in Ap values determined (in thepresence or absence of urea) would be expected to relateclosely to the urease activity of the cells, which is indeed thecase to some extent. The fact that Ap (in the presence ofurea) does not follow precisely the de novo ATP synthesismay suggest that energy is still required for housekeepingsystems at suboptimal pH values for growth. ATP synthesis,however, correlated to some extent with the differential inAp (in the presence or absence of urea; see below). Thepredominant component in the elevation of Ap values is Al,which contrasts with previous observations that ApH is themore significant factor (19, 21).These data extend a previous report that gave qualitative
information that the addition of urea to a suspension of U.urealyticum increased membrane potential (19). The in-crease in both Adj and ApH was totally abolished by theaddition of flurofamide, thus demonstrating that urea hydrol-ysis is, at a minimum, related to the increase in Ap.
Since we (13) and others have shown that the urease isentirely cytosolic and since only whole cells were used in ourshort assays, our studies show that cytosolic urea hydrolysisresulted in a concentration gradient of ammonia with aninternal concentration up to 21-fold greater than the externalconcentration. This suggests that passive diffusion of ammo-nia is limited. We have reported that some of the ammonia isutilized in citrulline synthesis (26). The most logical sourceof at least the increase in Ap is by conversion of the ammoniaconcentration difference into a chemical potential, i.e., theammonia chemical potential would equate approximately tothe whole of the AtJ and not just to the increase in A*observed in the presence of urea. Such a conversion wouldrequire that the membrane is permeable to ammonia, asNH4', via a saturatable uniporter (which would thus be arelatively slow process). It was noted that the externalammonia concentration remained constant despite variationof the internal ammonia concentration after the 30-s incuba-tion at each pH. The question then arises that if the mem-brane has a finite permeability for NH4' ions, why is anuncoupling cycle not set up with ammonia? This would beunlikely because the ratio of concentration of ammonia to
NH4+ ions will be very low in the assays, and, at therelatively acidic pH for optimum growth (25), to uncouple,ammonia would have to reenter the cell against a highconcentration gradient induced by urea hydrolysis. Althoughas the external pH increases, dissipation of the ammoniachemical potential by passive diffusion of ammonia couldincrease, such a situation should have a minimal effect(which cannot be calculated at present because the overallkinetics of the system remain to be clarified) at the externalpH values used in our assays.Although effective in abolishing electrochemical gradients
in other mycoplasmas (1, 7) and in our parallel studies withE. coli, several ionophores and uncouplers failed to affecteither Ad or ApH in U. urealyticum. There are severalpossible explanations for the inefficacy of these compounds.(i) The large flux of ammonia anticipated could swamp theireffect. They would not be expected to affect ammoniachemical potential, but they should abolish Ap (proton) byequating Ad and ApH terms to be equal and opposite. (ii)Ureaplasma membranes may be less susceptible as a resultof a peculiar (yet-unknown) structure associated with theorganisms being wall-less. (iii) A cation(s) other than protonsis involved in the energy transduction system. Or, (iv) thatthe ion-to-ATP stoichiometry is greater than that usuallyobserved. The answer to these questions is the basis offuture studies, particularly with other cations in combinationwith other ionophores.
Higher concentrations of CCCP were effective in givingsome reduction in Ad and in ATP synthesis, and this is inagreement with other data. Thus, other workers (19, 21)reported a 75% reduction in ATP synthesis in the presence of660 puM CCCP (a concentration at which CCCP may losespecificity and which could be inhibitory to other metabolicprocesses). This is consistent with only a small drop in AP,assuming that Ap is the driving force for ATP synthesis,since, as stated, the relationship is nonlinear.
In contrast to the relative inefficacy of ionophores, inhib-itors of both urease (flurofamide) and of FoF1 ATPase(DCCD) were effective in reducing ATP synthesis. Whenviewed along with the inhibition of ammonia chemical po-tential and the inhibition of the rise in Ap by flurofamide,inhibition of ATP synthesis indicates that hydrolysis of ureais the central factor in the synthesis of most ATP byureaplasmas and that the two processes are most likelycoupled by a chemiosmotic mechanism which may notconform exactly to the standard system. In fact, over the pHrange investigated, a closer correlation was observed be-tween the ammonia chemical potential, urease activity, andATP synthesis than between Ap (in the presence or absenceof urea), urease activity, and ATP synthesis. This furthersupports the interrelationship between urease activity andATP synthesis via ammonia chemical potential. Whether theATP synthesis is directed by end-product (NH4+) chemicalpotential or via a counter-ion-dependent ATPase remains tobe elucidated. The observed synthesis of ATP by the AT-Pase does not appear to be directly stimulated by ammoniumions alone. It has been shown (21) that NH4( ions do notstimulate ATP hydrolysis, and, in this study (data notshown), NH4' ions had no effect on ATP synthesis; thus,any mechanism would appear to be chemiosmotic. The FoF1ATPase inhibitor DCCD inhibited only 67% of ATP synthe-sis, and this may well be due to slow uptake and reaction ofDCCD during the relatively short preincubation period.Nevertheless, the inhibiting effect of DCCD appears toconfirm that ureaplasmas possess an ATP synthetase analo-gous to FoF1 ATPase.
The internal pH of ureaplasmas was always -0.85 pH unithigher than the external pH, in agreement with similarfindings for other mycoplasmas (1, 7). However, internal pHdoes not depend significantly on ammonia since, in thepresence of urea, only a small increase in internal pH wasseen. This small increase most likely occurs as a result ofscalar H' consumption as a consequence of urease action.The influence of external pH on ATP synthesis in urea-
plasmas has been reported (20), and the data presented hereare in broad agreement. In our study, it was noteworthy thatthe effects of exogenous urea, ionophores, and flurofamideon ATP synthesis mirrored their effects on the measurementof electrochemical parameters. Although exogenous or-thophosphate was included in the assays to potentiate ATPsynthesis, this was not necessary since ATP synthesis wasobserved in its absence (data not shown). This is despite areport (20) that its presence was essential. In addition, use ofphosphate buffer or of the standard buffer plus phosphateproduced values for electrochemical parameters similar tothose obtained with the standard buffer alone.Throughout the work presented, it is our contention that
any exogenous urea would be hydrolyzed within the shortassay periods used. We had previously reported (18) that atits optimum pH, purified ureaplasma urease is -100 timesmore active than jack bean urease. Assuming the knownurease content as a proportion of total cellular protein (12),the maximum amount of urea used in the assays would,under optimal conditions, be hydrolyzed in -7.5 s. Evenassuming suboptimal urea hydrolysis at the various pHvalues used, all added urea should be completely hydrolyzedwell within the 30-s assay period.With respect to ureaplasmas, it is of interest that the pH of
the urogenital tract is on the acid side of neutrality (29),similar to the pH values for optimum growth (25), maximumincrease in Ap, maximum ammonia chemical potential, max-imum urease activity, and maximum ATP synthesis. Thismay provide an explanation for the preferred host sites forcolonization.
Overall, the data presented indicate that urease activitygenerates an ammonia chemical potential with concomitantincreases in both Ap and ATP. Each of these were pHdependent and maximal at an external pH of 6.0, the optimalpH for ureaplasma growth. All were inhibited by flurofa-mide, supporting a link between these factors and confirmingthe hypothesis of Masover et al. (9) of a chemiosmoticmechanism for ATP generation in U. urealyticum resultingfrom urea hydrolysis. There are also indications that themetabolic processes of these simple organisms may be morecomplex than previously believed and that ureaplasmasdiffer significantly from other bacteria in this respect. How-ever, the nature of the cation(s) species involved, whether anammonia porter exists and whether the entire electrochem-ical potential generated during urea catabolism is due to anammonia chemical potential, will form the basis of futurework.
ACKNOWLEDGMENTS
This work was supported by a grant (G8924533) from the MedicalResearch Council.
I. Armitt and P. McCready are thanked for technical assistance.
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