JOURNAL OF BACTERIOLOGY, Feb. 1987, p. 823-829 Vol. 169, No. 20021-9193/87/020823-07$02.00/0Copyright © 1987, American Society for Microbiology

In Vivo Regulation of Histidine Ammonia-Lyase Activity fromStreptomyces griseus

TERRY A. KROENING AND KATHLEEN E. KENDRICK*Department of Microbiology, The Ohio State University, Columbus, Ohio 43210

Received 10 November 1986/Accepted 21 November 1986

The enzyme histidine ammonia-lyase (histidase) is required for growth of Streptomyces griseus on L-histidineas the sole source of nitrogen. Histidase was induced by the inclusion of histidine in the medium, regardless ofthe presence of other carbon and nitrogen sources. Histidase activity was increased by a shift of cultureincubation temperature from 30 to 37°C. Conversely, upon induction of sporulation by either phosphatestarvation or nutritional downshift, histidase underwent rapid inactivation. Nutrient replenishment fullyreversed histidase inactivation while simultaneously permitting reinitiation of vegetative growth. In contrast tohistidase inactivation during sporulation, histidase was activated after transition of a vegetatively growingculture to stationary phase. Although neither activation nor inactivation required de novo protein synthesis,inactivation appeared to involve a heat-labile protein. The results indicate that histidase activity is regulated invivo by a process that responds to changes in the growth phase of the organism.

To utilize L-histidine as the sole source of nitrogen,streptomycetes require the enzyme histidine ammonia-lyase(histidase; EC 4.3.1.3; 8). This enzyme catalyzes thenonoxidative deamination of L-histidine, forming urocanateand ammonia. In bacteria capable of utilizing histidine as acarbon source, catabolism of urocanate results in the gener-ation of glutamate, which can then be used in biosynthesis oras a source of energy and reducing power (6, 12, 13, 16, 23).

In the enteric bacteria (1, 16, 22), pseudomonads (11, 12,19), and Bacillus subtilis (2, 5), histidase synthesis is inducedwhen histidine is the sole carbon and nitrogen source,whereas histidase is not synthesized in the absence ofexogenous histidine. Histidase expression is not guaranteedby the presence of histidine in the environment, however. InAerobacter aerogenes histidase gene expression is subjectnot only to induction but also to carbon catabolite repressionand nitrogen regulation; histidase synthesis is repressed byglucose, but only in the presence of a readily utilizablenitrogen source such as ammonia (18, 20). Contrastingresults were obtained in the analysis of the regulation ofhistidase expression in B. subtilis and Salmonellatyphimurium. Investigators observed induction as well ascarbon-catabolite repression of histidase; however, histidasegene expression in these organisms does not occur undernitrogen-limiting conditions in the presence of glucose (1, 2).Our previous results (8) suggested that histidase from the

gram-positive organism Streptomyces coelicolor is subject toneither carbon catabolite repression by glucose nor nitrogenregulation by ammonia. As in other bacteria, however,histidase is inducible in streptomycetes. In this paper, weshow that the presence of L-histidine in both defined andcomplex media induces histidase in Streptomyces griseus,regardless of the presence of additional carbon and nitrogensources. Furthermore, kinetic analysis of histidase from S.griseus suggests that histidase activity is regulated in vivo bya novel, reversible posttranslational mechanism.

(These results were presented in part at the 86th AnnualMeeting of the American Society for Microbiology [T. A.

* Corresponding author.

Kroening and K. E. Kendrick, Abstr. Annu. Meet. Am.Soc. Microbiol., 1986, K-60, p. 203].)

MATERIALS AND METHODSGrowth of bacterial strains. For all experiments, S. griseus

strain NRRL B-2682 was used. An activated submergedspore preparation of strain 2682 (7), pregerminated in Tryp-ticase soy broth (BBL Microbiology Systems, Cockeysville,Md.), was inoculated into the appropriate medium to aconcentration of approximately 5 x 105 CFU/ml and grownovernight at 30°C in a rotary shaker at 250 rpm. Polyethyleneglycol 8000 (5% [wt/vol]; Sigma Chemical Co., St. Louis,Mo.) and a coiled spring were included in all media toenhance dispersed growth (8). Medium 2XYT was preparedaccording to the formula of Miller (17). When 10 mML-histidine was added to the 2XYT (designated 2XYTH), 10mM N-tris(hydroxymethyl)methyl-2-aminoethanesulfonicacid (TES)-NaOH or sodium-potassium phosphate (NaKP1)buffer, pH 7.3, was also included. The phosphate buffer wasprepared as a 1 M stock solution by combining 1 M solutionsof NaH2PO4 and K2HPO4 to obtain the appropriate pH.Sporulation medium (SM; 7) contained 1.0 mM K2SO4, 0.1mM FeCl3, 0.1 mM sodium nitrilotriacetate, 50 mM NaKP,(pH 7.3), 2 mM MgCl2, 0.1 mM CaCI2, 20 mM glucose, 20mM NH4Cl, and 10 mM L-histidine (added as a filter-sterilized solution of 0.5 M L-histidine-NaOH, pH 6.0). Toinduce sporulation by phosphate starvation, Pi was omittedfrom this medium and 50 mM TES-NaOH (Sigma), pH 7.3,was included. When sporulation was induced by nutritionaldownshift, 50 mM NaKP1, pH 7.3, was used as buffer. Othermedia formulations and conditions of growth were as de-scribed by Kendrick and Ensign (7), with the addition of 10mM L-histidine when needed. The complex media used inthis study, 2XYT and SM supplemented with 1% (wt/vol)casein hydrolysate (vitamin-free, salt-free; ICN NutritionalBiochemicals, Cleveland, Ohio) (SM + CAA), prohibitedsporulation of strain 2682.When sporulating cultures were to be assayed for

histidase, induction of sporulation was effected by the fol-lowing method. A vegetatively growing culture (250 ml, in

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824 KROENING AND KENDRICK

o 0.9f-

:1

5TIME (min)

i.7

1.6

1.5 °'

1.4 ,<

1.3

1.2

FIG. 1. Histidase reaction profile. A crude extract prepared from vegetative mycelia grown in 2XYTH was assayed for histidase at 30°Cby measuring the appearance of product, urocanate, which absorbs at 277 nm. Reaction conditions were as specified in the text. Symbols:O, unheated extract; 0, extract heated at 60°C for 10 min before the assay.

phosphate-buffered SM + CAA) was harvested at roomtemperature by centrifugation at 15,000 x g for 10 min. Themycelial pellet was washed with 30 ml of SM and centrifugedagain. The cell pellet was then suspended in 10 ml of SMprewarmed to 30°C, and this suspension was rapidly trans-ferred to 250 ml of identical medium in a 1-liter Erlenmeyerflask. This entire process was completed in approximately 40min. The time at which the mycelia were added to the flaskwas designated 0 h.Enzyme analysis. The cells and materials used in the

preparation of the extract were maintained at 4°C throughoutthe procedure. Mycelia were harvested by centrifugation at27,000 x g for 15 min. Cells were washed with 1 M KCl andconcentrated 50-fold in 50 mM Tris hydrochloride, pH 7.5.After cell disruption by a single passage through a Frenchpressure cell at 12,000 lb/in2, cell debris was removed bycentrifugation at 27,000 x g for 15 min. The supematant atthis stage served as the crude extract. In some cases, thecrude extract was subsequently centrifuged at 100,000 x gfor 1 h to remove particulate material, and the resultingsupernatant was dialyzed against 200 volumes of extractionbuffer for a total of 2 h (one change) before the assay. Thissource of enzyme was designated dialyzed extract. As analternative to dialysis, the supematant was passed through aPD-10 desalting column (Pharmacia Fine Chemicals, Piscat-away, N.J.).The assay for histidase activity was that used by Kendrick

and Wheelis (8). One ml of 50 mM Tris hydrochloride, pH8.5, was prewarmed to the reaction temperature. Approxi-mately 150 ,ug of protein (25 ,ud of cell extract) was added,immediately followed by 50 ,ul of 0.1 M L-histidine-NaOH(pH 6.0; Calbiochem-Behring, La Jolla, Calif.) to start thereaction. The reaction was monitored in a thermally regu-lated recording spectrophotometer (Gilford Instrument Lab-oratories, Inc., Oberlin, Ohio) for at least 10 min. Unlessotherwise indicated, assays were conducted at 30°C. Specificactivity was calculated as nanomoles of urocanate formedper minute (milliunits) per milligram of protein, with 18.8cm-l pmol-1 ml as the millimolar extinction coefficient for

urocanate (8). Protein concentration was measured accord-ing to the spectrophotometric method of Ehresmann et al.(4), with bovine serum albumin as a standard. For a givenextract preparation, the initial reaction velocity was linearlyproportional to the concentration of protein.For heat treatment, a sample of the extract was added to

a glass tube and incubated at the indicated temperature forthe duration specified in Results. For most experiments, atemperature of 60°C was used. In the heat-activation exper-iment, however, samples of the extract were incubated attemperatures between 4 and 80°C.

Activation rate ("Activation"; see Tables 1, 2, and 3) wascalculated as the ratio of activity at 9.5 min of reaction toinitial activity, measured 1.0 min after initiation of thereaction. Each activity was calculated by measuring thechange in absorbance during a 1-min period, from 9.0 to 10.0min in the former case and from 0.5 to 1.5 min in the lattercase. The value obtained for activation represents the rate ofconversion of inactive histidase to active histidase. Percentactive ("% Active"; see Tables 2 and 3) refers to the fractionof histidase existing in the active form and was determinedas 100 x the initial activity at 1.0 min of reaction relative tothe activity in the extract heated at 60°C for 10 min.

RESULTS

Histidase activity is inducible in Streptomyces griseus.Growth rate measurements revealed that histidine is anexcellent nitrogen source for S. griseus. In the absence ofL-histidine, virtually no histidase activity was detected. Thespecific activity of histidase increased approximately 50-foldupon inclusion of histidine in the medium of exponentiallygrowing cultures, regardless of the presence of glucose,glutamate, or ammonium chloride.

Histidase undergoes activation in vitro. In the course ofanalyzing histidase activity in crude extracts prepared fromvegetative mycelia of S. griseus, we observed that thereaction kinetics displayed a curvature indicative of anactivation event (Fig. 1). We quantified this activation as the

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ratio of activity at 9.5 min of reaction to the initial activity,measured at 1.0 min of reaction. When histidase fromexponentially growing cultures was assayed in crude ex-tracts at 30°C, activation approximated 2.0. Histidase acti-vation was probably not caused by the presence of substrateor product: incubation of the reaction mixture, containingdialyzed extract, at 30°C for 10 min in the absence ofL-histidine activated the enzyme to the same extent asincubation in the presence of L-histidine (Table 1). A varietyof other small molecules likewise had no effect on theactivation of histidase. Unlike pseudomonad histidase,which is activated by thiol reagents and manganous ion (9,14, 15, 21), streptomycete histidase activity was altered byneither thiol reagents nor divalent cations nor EDTA (Table1).The activation observed during the histidase reaction

could be interpreted to mean that multiple forms of histidaseexist in the extract. To investigate this possibility, wemeasured the histidase reaction rate at a variety of temper-atures. Figure 2 shows an Arrhenius plot for untreated crudeextract. The optimum reaction temperature for streptomy-cete histidase was approximately 50°C. More informative,however, was the discontinuity in the slope of this plotbetween 30 and 45°C. Linear regression analysis of thesedata indicated two distinct lines that intersect at 36.9°C, withslopes of -1.3 and -3.5. This result is consistent with theoccurrence of two forms of histidase in the crude extract,each catalyzing reactions with distinct energies of activation.The curved kinetics of the histidase reaction prompted us

to determine whether histidase in crude extracts could beactivated by heat treatment in vitro. Identical portions ofcrude extract prepared from a culture of strain 2682 grown tolate exponential phase in 2XYTH were heated for 10 min attemperatures from 30 to 80°C. Partial activation occurredduring incubation of the extract for 10 min at 30°C. Treat-ment at temperatures between 40 and 70°C resulted inuniformly high activity with linear reaction kinetics (Fig. 1),whereas histidase rapidly lost activity when the crude ex-tract was heated to 80°C. The activation was not reversed bysubsequent prolonged incubation at 4°C, nor was the in-

TABLE 1. Effect of substrate and cations on histidase activity indialyzed crude extract

Addition to Preincubationb Sp act at 30'CC Activationreaction' (mM) at 30°C (min) 1.0 min 9.5 min

None 0 6.7 ± 0.4 13.2 ± 0.6 2.010 13.0 ± 0.3 13.8 ± 0.3 1.1

BMEd (1.3) 0 6.3 ± 0.5 13.0 ± 0.6 2.110 12.5 ± 0.4 14.0 ± 0.5 1.1

DTTI (1.0) 0 6.5 ± 0.5 13.5 ± 0.6 2.110 12.6 ± 0.4 14.0 ± 0.2 1.1

EDTA (0.1) 0 6.6 ± 0.5 12.9 ± 0.5 2.010 12.8 ± 0.4 14.1 ± 0.3 1.1

MnCI{f (0.1) 0 6.7 ± 0.6 13.2 ± 0.6 2.0a The standard reaction mixture contained 47 mM Tris hydrochloride (pH

8.5), 4.7mM L-histidine, and 100 to 200 jig of protein in a final volume of 1.075ml.

b The reactions were initiated by the addition of 5 mM L-histidine, eitherimmediately (0 min) or after preincubation of the incomplete reaction mix,lacking histidine, at 30°C (10 min).

c In milliunits per milligram of protein; results are expressed as mean +standard deviation of six assays.

d Identical results were obtained when the concentration of P-mercaptoeth-anol (BME) was reduced to 1.0 mM or increased to 5.0 mM.eDTT, Dithiothreitol.f Essentially identical results were obtained when CdCl2, MgCl2, CaC12,

ZnCl2, or CoCl2 was included in the reaction mixture at 0.1 mM.

TEMPERATURE (0C)60 55

1.50, F

>- 1.40

H>

u 1.30LL

&uII

0-O 1.20-J

50 45 40 35 30

o 3.10 3.20TEMPERATURE' (10-3.OK-')

3.30

FIG. 2. Arrhenius plots of histidase activity. A crude extract wasprepared from a culture grown in 2XYTH to late exponential phase.One half of the extract was held at 4°C before the assay (A), whereasthe remainder of the extract was heated at 60°C for 10 min (O) beforebeing assayed at the indicated temperatures. Each point is theaverage of assays conducted in triplicate; for no point did thestandard deviation exceed 1.6% of the average. The slopes of thelines, determined by linear regression analysis, were -3.5 (line 1),-1.3 (line 2), and -1.2 (line 3). Lines 1 and 2 intersect at 36.9°C.

creased activity caused by a decrease in the protein concen-tration of the heated extracts.We then compared the kinetics of histidase activation

during incubation of the crude extract at three temperatures.Samples of a crude extract prepared from mycelia grown tomid-exponential phase in SM + CAA were incubated at 30,37, or 60°C. At various times, samples were removed andchilled at 4°C. After all samples had been collected, theywere assayed for histidase at 30°C. Activation was mostrapid at 60°C; histidase acquired maximal activity after 7 minof incubation (Fig. 3). Activation was complete in 10 min at37°C, whereas full activation at 30°C required incubation ofthe extract for 40 min. The activation kinetics in the reactionmixture closely paralleled that of the undiluted extractincubated at 30°C: 30 min of reaction was required to obtainfully active histidase, and the maximal activity attained wasidentical to that reached after incubation of the undilutedextract at 30°C for 40 min (data not shown).The Arrhenius plot for extract heated at 60°C for 10 min

suggested the presence of a single enzyme form with an

optimal reaction temperature of 45 to 48°C (Fig. 2). Theslope of the line for the heated extract (-1.2), as determinedby linear regression analysis, was essentially identical to thatobtained for the unheated extract when assayed at temper-atures higher than 36°C (-1.3).

If the temperature of discontinuity of the unheated extract(37°C; Fig. 2) were physiologically significant, we wouldexpect that histidase in mycelia grown at relatively hightemperature would exist primarily as the active form. To testthis prediction, strain 2682 was grown at 30°C in SM + CAA

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'::__' 25

0

tL20-

pE

UT

E

10 20 30 40 50 60INCUBATION (min)

FIG. 3. Kinetics of heat-induced activation of histidase. Samplesof crude extract, prepared from vegetative mycelia grown in SM +CAA, were incubated at 30 (0), 37 (A), or 600C (O). At the indicatedtimes, samples were withdrawn and rapidly chilled at 4°C forapproximately 2 h. Subsequently, all samples were assayed forhistidase activity at 30°C. Data are presented as the average of atleast six assays.

to an A420 of 1.0. Equal portions of this exponentiallygrowing culture were incubated for an additional 30 min at 30and 37°C, during which time both cultures continued to growat approximately equal rates. Crude extracts were thenprepared and assayed for histidase. Growth of the mycelia at370C resulted in virtually all of the histidase existing in itsactive form (99% active) displaying linear reaction kinetics,whereas in the mycelia grown at 30°C, histidase was main-tained in a less active form (39% active) with a moderate rateof activation (Table 2). The presence of chloramphenicol at200 ,ug per ml, a concentration sufficient to inhibit at least85% of protein synthesis in strain 2682 (K. E.K., unpublishedobservations), did not affect the in vivo activation ofhistidase (Table 2).The results of our in vitro studies were consistent with the

hypothesis that the conversion of active histidase to the lessactive form was sensitive to high temperature. To determinewhether histidase inactivation was heat sensitive in vivo, weattempted to reverse the heat-induced activation of histidaseby transfer of the mycelia from 37 to 300C. In the absence ofchloramphenicol, histidase was partially inactivated upontransfer from 37 to 300C (Table 2). When protein synthesiswas inhibited by the inclusion of chloramphenicol, however,histidase was maintained in a fully active form (Table 2).

Histidase is inactivated during sporulation. Our resultsindicated that streptomycete histidase activity was modu-lated in vivo by changes in the growth temperature. Wewished to learn whether other culture incubation conditionsalso affected histidase activity. To this end, extracts wereprepared from cultures growing vegetatively as well as fromcultures that had been induced to sporulate by phosphatestarvation for 15 min, or 1 or 2 h. Histidase activity andactivation rate were compared in these four extracts.Histidase in the vegetatively growing culture remained at anintermediate activity (31% active), with an activation rate of2.0 (Table 3). In contrast, histidase activity rapidly de-creased during the earliest period of sporulation (Table 3), atwhich time histidase existed primarily as the less active form

(18% active). The fraction of active histidase graduallyincreased as sporulation proceeded. In all of the sporulatingcultures, the activation rate was high. (The difference inactivity between the vegetative culture and the sporulatingculture is significant by Student's t test analysis [P < 0.001]).The activities measured after heat treatment of the extractsindicated that the total amount of histidase present under allculture conditions was approximately equal (P > 0.1).We then tested the possibility that the inactivation of

histidase in a sporulating culture was a consequence ofphosphate starvation rather than the induction of sporula-tion. To do this, we induced sporulation by nutritionaldownshift. The kinetics of the sporulation process inducedby nutritional downshift are essentially identical to those ofsporulation during phosphate starvation (7; unpublishedobservations). When exponentially growing mycelia weretransferred from a complex medium (SM + CAA) to SM(from which the casein hydrolysate was omitted), a crudeextract prepared from mycelia harvested 15 min after trans-fer contained histidase that was 12% active (Table 3), with anactivation rate of 4.2. The fraction of active histidase in-creased twofold during the course of the experiment, and theactivation rate remained high (Table 3).To demonstrate that the rapid decrease in histidase activ-

ity was not the result of the mycelia entering nongrowingconditions, we also measured histidase activity in an extractprepared from stationary-phase mycelia. A greater propor-tion of histidase from this culture existed as the more activeform (57% active; Table 3).

Histidase inactivation is reversible. Vegetatively growingcultures were induced to sporulate by the nutritionaldownshift procedure described above. At 15 min after induc-tion was initiated, casein hydrolysate was added back to theculture to a final concentration of 1%. No additions weremade to the control culture. After 15 min or 1 h of furtherincubation, mycelia were harvested, disrupted, and assayedfor histidase. The results indicate that the nutritionallyreplenished culture, which had resumed growth, containedhistidase that was significantly more active (P < 0.001) thanthat of the sporulating culture (Table 3). Similar results wereobtained when chloramphenicol was added to the mycelia 15min before the initiation of the nutritional downshift (Table

TABLE 2. Effect of growth temperature on histidase activityTemp Initial sp actb(°C) of Addition Unheated Heated Active' Activation

incubation" extract extract'

30 None 8.3 ± 0.2 21.3 ± 0.9 39 2.0 ± 0.0CAMe 6.7 ± 0.2 20.2 ± 0.9 33 2.1 ± 0.0

37 None 38.5 ± 0.7 39.0 ± 0.4 99 1.0 ± 0.0CAM 35.7 ± 0.3 38.1 ± 0.8 94 1.0 ± 0.0

37, then 30f None 17.9 ± 0.3 22.9 ± 0.5 78 1.2 ± 0.0CAM 31.1 ± 0.7 32.8 ± 1.2 95 1.1 ± 0.0

a A culture was grown at 300C in SM + CAA to an A420 of 1.0, at whichpoint equal portions were rapidly equilibrated to the indicated temperatureand incubated for 30 min. Mycelia were then harvested and assayed forhistidase at 30°C.

b In milliunits per milligram of protein; results are expressed as mean ±standard deviation of six assays.

Extract heated at 60°C for 10 min.d Percent activity in the unheated extract relative to that in the heated

extract.e Chloramphenicol (CAM) added at 200 ,ug ml-', 15 min before division of

the culture.f A culture was grown at 30°C, then transferred to 37°C for 30 min.

Subsequently, the culture was transferred back to 30°C for another 30 min.The mycelia were then harvested and assayed by the standard technique.

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STREPTOMYCETE HISTIDASE 827

TABLE 3. Effect of growth condition on histidase activityInitial sp actc

Culture condition Duration (min)a No. of trialsb Unheated Heated % Active' Activationextract extractd

GrowingVegetative NA 12 7.2 ± 1.3 23.1 ± 1.5 31 2.0 ± 0.1Nutritionally replenished 15 3 8.8 ± 1.5 20.8 ± 1.3 42 1.6 ± 0.1

60 1 12.9 ± 0.5 23.5 ± 0.7 55 1.2 + 0.0+ CAMf 15 3 13.6 0.6 24.2 ± 0.5 56 1.7 ± 0.1

SporulatingPhosphate-starved 15 3 3.9 + 1.1 22.2 ± 1.1 18 3.0 ± 0.0

60 4 5.3 + 0.6 24.5 ± 0.8 22 3.1 ± 0.1120 4 6.4 ± 0.4 23.2 ± 0.9 27 2.9 ± 0.1

Nutritionally downshifted 15 4 2.5 ± 0.2 21.6 ± 0.7 12 4.2 ± 0.230 3 4.7 ± 0.2 23.2 ± 0.5 20 3.1 + 0.160 3 4.1 ± 0.4 21.3 ± 0.8 19 3.1 ± 0.2120 3 4.7 + 1.0 23.1 ± 1.4 20 3.1 ± 0.1

+ CAM 30 3 5.7 ± 0.4 23.9 ± 0.4 24 3.0 ± 0.1

Stationary phase NA 3 12.0 ± 0.9 21.2 ± 1.5 57 1.2 ± 0.1a Time at the specified culture condition. NA, Not applicable.b For each trial, assays were performed in triplicate.In milliunits per milligram of protein; results are expressed as mean ± standard deviation.

d Extract heated at 60'C for 10 min.e See Table 2, footnote d.f When used, chloramphenicol (CAM; 200 p.g ml-) was added to the cultures 15 min before the induction of sporulation and was allowed to remain for the rest

of the incubation period.

3), indicating that the reversibility was independent of pro-

tein synthesis during continuous incubation at 30°C.

DISCUSSION

We conclude from these experiments that streptomycetehistidase expression and activity appear to be regulated bydistinctive mechanisms. Histidase was induced wheneverhistidine was included in the culture medium, regardless ofthe presence of other carbon and nitrogen sources. Thisobservation is not surprising in view of the fact that histidineis an excellent nitrogen source for S. griseus, and theobservation agrees with the results obtained in the analysisof histidine utilization in S. coelicolor (8). It appears, there-fore, that histidase expression in streptotnycetes is notsubject to either carbon catabolite repression by glucose orglutamate or nitrogen regulation by ammonia or glutamate.Our results also demonstrate that histidase activity is

regulated in vivo. Initial observations indicated thathistidase from both S. griseus and S. coelicolor underwentactivation during the enzymatic reaction; additionally,histidase activity could be maximized and made linear byheat treatment (Fig. 1; K. E. Kendrick, Ph.D. thesis, Uni-versity of California, Davis, 1979). Because heat treatmentresulted in an increase in activity of the extract whenassayed at 30°C, we conclude that heat treatment convertsthe less active form of histidase to the more active form invitro. The active form generated by treatment of the extractat 60'C for 10 min is likely to be identical to that generatedduring the histidase reaction, since both conditions resultedin enzymatic reactions with the same energy of activation(Fig. 2).The temperature shift experiment (Table 2) showed that

the in vitro activation process directly correlates with in vivoactivation. Histidase inactivation appears to involve a heat-sensitive event which is dependent on one or more proteins(Table 2). The simplest interpretation of this result is thathistidase inactivation requires the participation of a heat-

labile protein. Additional experiments suggested thathistidase inactivation may be characteristic of the initialstage of streptomycete sporulation, whether induced byphosphorus starvation or nutritional downshift (Table 3). Wehave therefore identified four culture conditions which havedistinct effects on histidase activity: vegetative growth atlow temperature, growth at high temperature, entry intostationary phase, and sporulation. We note that incubationat 37°C, which results in activated histidase, inhibits sporula-tion of S. griseus (K. E. Kendrick, unpublished observa-tions). Histidase inactivation at the onset of sporulation isreversed by conditions that allow mycelia to renew vegeta-tive growth. Furthermore, both activation and inactivationare independent of protein synthesis in cells incubated at30°C (Table 3).There are three possible mechanisms by which histidase

activity could be modulated both in vitro and in vivo.Perhaps histidase can undergo a reversible change in con-formation dependent on the culture conditions. McClard andKolenbrander (14, 15) have shown that histidase purifiedfrom Pseudomonasfluorescens comprises two conformation-al isomers which can be interconverted by temperature orinteraction with sulfhydryl reagents. These researchers alsodemonstrated a discontinuous Arrhenius plot for purifiedhistidase (14, 15). No evidence was presented, however, toindicate that this mechanism regulated histidase activity invivo.

Alternatively, histidase inactivation may occur by revers-ible polymerization, as Klee (9) has suggested for histidasepurified from pseudomonad strain 11299. The polymerizedforms of histidase were less active than the monomeric form.In agreement with the data of Rechler (21), Klee (9) detectedsulfhydryl-dependent and cation-dependent activation of pu-rified histidase ahd consequently proposed that depolymer-ization required interaction of histidase with sulfhydrylreagents. Consevage and Phillips (3), however, were unableto confirm the existence of histidase polymers in theirpreparation of histidase from Pseudomonas putida. In con-

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trast to the results obtained with pseudomonad histidase, wehave been unable to demonstrate a dependence on eithersulfhydryl reagents or divalent metal cations for the activa-tion of histidase from either S. griseus (Table 1) or S.coelicolor (Kehdrick, Ph.D. thesis). Our results suggest thatthe activation of streptomnycete histidase occurs by a mech-anism that is distinct from the sulfhydryl-induced activationof purified pseudomonad histidase.

Direct interaction with another metabolite present in themycelia also could cause the reversible activation ofhistidase. In light of the heat sensitivity of inactivation(Table 2), we think that this third mechanism would be morelikely to account for modulation of histidase activity in vivo.It is difficult to conceive of an enzyme which, upon treat-ment at moderate temprature, retains full catalytic activitywhile losing all self-regulating properties. Because dialyzedcrude extract retained the ability to be activated, either theactivating factor is relatively large (Mr> 10,000) or activa-tion is a consequence of removal of a factor from histidase.We have obtained identical results when the crude extractwas passed over a gel filtration column with an exclusionlimit of 5,000 daltons (data not shown). This result rules outactivation of histidase via interaction with a cofactor. Acti-vation processes such as dissociation of a tightly boundinhibitor, or covalent modification of histidase, are certainlypossible. Our data appear to exclude proteolysis as theinactivation mechanism, as histidase was rapidly activatedduring a period of reduced protein synthesis (Tables 2 and 3).Furthermore, the in vitro kinetics, displaying activation, are

inconsistent with a proteolytic-inactivation mechanism.Activation of histidase by proteolysis is more difficult to

rule out. If proteolysis of the less active enzyme generatedthe active form, then we would expect activation to beirreversible. Our results indicate that activation byproteolysis is unlikely because inactivation occurred rapidly,independent of protein synthesis, during the induction ofsporulation (Table 3). Because proteins were still beingsynthesized, albeit at a low rate, it is alternatively possiblethat the production of less active histidase upon induction ofsporulation is a consequence of leaky translation of thehistidase message in the presence of chloramphenicol. Thetemperature effects in vivo, however, strongly suggest thatthere is at least one protein involved in histidase inactivation(Table 2); this result is inconsistent with a proteolyticactivation mechanism, for which there would be no conver-sion to the less active form. The most straightforwardinterpretation of the data presented here is that histidaseactivity is regulated posttranslationally by a protein-dependent mechanism that converts active histidase to aninactive form.Even though histidase inactivAtion at the onset of sporula-

tion involves only a two- to threefold decrease in activity,this subtle change in activity may reflect a profound changein cellular metabolism. LaPorte et al. (10) have demon-strated that a fourfold increase in the activity of isocitratedehydrogenase from E. coli, effected by the addition ofglucose to a culture growing on acetate, is amplified into a97-fold increase in the flux of isocitrate through the citricacid cycle. As a consequence, flux through the glyoxylateshunt is virtually eliminated (10). The authors presentedtheoretical and empirical arguments that minor alterations ineither Vmax or Km can drastically alter cell metabolism at ametabolic branch point (10). Histidine does occur at a branchpoint in streptomycete metabolism. Conversion, viahistidase and subsequent enzymes, to glutamate is necessaryfor utilization of histidine as a carbon and nitrogen source for

vegetative growth, whereas it is unlikely that histidase isrequired for sporulation, since S. griseus sporulates well indefined media in the absence of exogenous histidine (7). Wespeculate that it is advantageous to the streptomycete toincrease the flux of histidine through protein synthesis, andperhaps also production of secondary metabolites, at theonset of sporulation. Therefore, a minor decrease inhistidase activity upon exposure to conditions that inducesporulation may have the potential to redirect histidinemetabolism toward sporulation-specific processes.Although our results implicate a reversible inactivation

event in the regulation of histidase activity, we cannot yetdiscriminate among the three basic mechanisms by whichthis inactivation may occur. We are currently purifyinghistidase and analyzing mutants defective in histidine utili-zation to determine the biochemical mechanism underlyinghistidase inactivation during streptomycete sporulation.

ACKNOWL1EDGMENTSWe thank P. S. Perlman and C. W. Birky, Jr., for the use of their

spectrophotometer. We are also grateful to J. C. Ensign; M. T.Muller, J. N. Reeve, W. R. Strohl, and R. P. Swenson for criticallyreviewing the manuscript.

This work was supported by a grant from the Ohio State Univer-sity Office of Research and Graduate Studies.

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