The relationship of serine protease activity to RNA polymerase modification and sporulation in Bacillus subtilis

J. Mol. Biol.

(1973) 76, 103-122

The Relationship of Serine Protease Activity to RN Polymerase Modification and Sporulation in Bacillus subtilis T. J. LEIGHTON~, R. H. DOI Department of Biochemistry and Biophysics University of California, Davis, Calij,, U.S.A. R. A. J. WARREN AND R. A. KELLN Department

of Microbiology,

Vancouver,

University of British B.C., Canada

Columbia

(Received 17 August 1972, and in revised Sorun 12 JorLuary 1973) The isolation and properties of a single site temperature sensitive protease mutant of Bacillus subtilis are described. Numerous criteria suggest that the mutation resides in the structural gene coding for a basic serine protease. The mutation has been mapped between aroD and lys-1 on the Bacillus subtilis chromosome. This protease exists as an intracellular and extracellular enzyme. The mutant cells are temperature sensitive for sporulation, antibiotic production, and the sporulation-specific alteration in DNA-dependent RNA polymerase p subunit. Several of this enzyme in a limited proteotypes of evidence indicate a direct involvement lytic cleavage of vegetative RNA polymerase /3subunit, which produces the lower molecular weight ,8 subunit found in sporulating cells. The derangement in this process is sufficient to account for the stoppage of sporulation at stage 0 when the mutant cells are grown at the non-permissive temperature.

1. Introduction The relationship between the post-exponential appearance of extracellular proteolytis enzymes and bacterial sporulatiou has been studied by many investigators (Spizizen, 1965; Schaeffer, 1969; Aronson et al., 1971; Sadoff et al., 1970; Michel & Millet, 1970; Millet, 1969,197O; Millet & Aubert, 1969; Hoch & Spizizen, 1969; Balassa, 1969; Prestidge et al., 1971; Bernlohr & Clark, 1971). There has been no general agreement as to whether these enzymes are functionally or fortuitously related to spore production. Recent detailed biochemical analysis suggests that several pmteases are found in the culture supernatant fraction of sporulating Bacilli and that the number and relative amounts of these enzymes vary widely between the strains of Bacilli studied. Hence, in Bacillus sub&s most of the extracellular proteolytic activity directed against casein is due to a basicf sorine protease (Millet, 1970; Preatidgc et al., 1971; Boyer & Carlton, 1968; Hageman & Carlton, 1970). The culture supernatant fraction also contains a basic metal protease and an acidic serine esterase $ Present address where part of this work was carried out: Department University of Massachusetts Medical School, 419 Belmont Street, Worcester, $ The terms basic and acidic are used here with reference to the isoelectric 103

of Microbiology,

Mass. 10604, U.S.A. point of t’he protein.

104

T. J. LEIGHTON

E!Z’ AZ.

(Boyer & Carlton, 1968; Millet, 1970; Prestidge et al., 1971). In the case of B. megaand B. cereus most of the extracellular proteolytic activity is due to a metal protease (Aronson et al., 1971; Millet, 1969). B. cereus has been shown also to produce low levels of a serine protease (Sadoff et al., 1970). It is clear from these data that a meaningful comparison of mutants which cannot hydrolyze casein and/or sporulate is not possible between these species of Bacilli since different enzymes may be affected. In addition there has been no compelling demonstration that any of the mutants analyzed affect the structural gene required for proteolytic activity. In fact the primary characteristic of most of the protease mutants examined is that the levels of several of these enzymes are concomitantly decreased (Schaeffer, 1969; Michel & Millet, 1970). Also, the intracellular protease activity in mutant cells has not been examined. There is circumstantial evidence that the metal protease may not be required for sporulation in B. s&&s, B. megaterium and B. cereus (Aronson et al., 1971; Michel & Millet, 1970; Millet & Aubert, 1969). We have previously demonstrated that a single-site temperature sensitive basic serine protease mutant (ts-5) can block sporulation biochemically, physiologically and morphologically at stage 0 (Leighton et al., 1972; Santo et al., 1972). When ts-5 cells are grown at the non-permissive temperature the sporulation-specific change in RNA polymerase template specificity and subunit composition (Losick & Sonenshein, 1969; Losick, 1970) does not occur. Therefore, the basic serine protease is required for the initiation of sporulation in B. subtilis. In this communication we present further biochemical and genetic characterization of this mutant. terium

2. Materials and Methods (a) Bacterial &rains ati viruses WB746 (wild type B. subtilis Marburg) was obtained from Dr E. Nester. B. subtilis Br16 (lys-, trp-), B. subtilis SB120 (are-, trp-), B. subtilis JH305 (strC2, trp-), and B. subtilis strain H were from Dr J. Hoch. Bacteriophage PBS1 was from Dr F. E. Young and bacteriophage

4e was from

Dr R. Losick. (b) Chemicals and supplies

Tryptose blood agar base, Penassay

medium and nutrient broth (Difco Labs, Detroit, Michigan). UTP, GTP, CTP and ATP (Boehringer Mannheim, New York). Phenyl methane sulfonyl fluoride, dithiothreitol, bovine serum albumin, chloramphenicol and azocasein (Cal Biochem, San Diego, Calif.). Tris, ammonium sulfate, urea and Coomassie Brilliant Blue (Mann Research Labs, New York). Ethane methyl sulfonate and glycerol (Eastman Organic Chemicals, Rochester, N.Y.). Aorylamide and N,N’-methylene-bisacrylamide (Bio-Rad, Richmond, Calif.). 0-Phenanthroline, N,N,N’,N’-tetramethylethylenediamine and ammonium persulfate (Matheson Coleman and Bell, East Rutherford, N.J.). Sodium dodecyl sulfate (Sigma Chemical Co., St. Louis, MO.). DEAESephadex, A50; CM-cellulose Sephadex, C50 and G75 Sephadex (Pharmacia, Piscataway, Elkhart, Indiana). Fiber-glass filters, AP200N.J.). Poly(dA-dT) (Miles Laboratories, 2700, and 13 mm membrane filters, HAWPO1300 (Millipore Corp., Bedford, Mass.). [3H]UTP (New England Nuclear, Boston, Mass.). (c) Growth

conditions

All strains were maintained as isolated colonies on Tryptose agar base) incubated at room temperature. Bacteria were grown nutrient broth containing O.1o/o glucose (Leighton & Doi, 1971). transferring a 2-day old colony into growth medium. When the units (Klett Summerson Calorimeter, no. 66 filter) a 5% (v/v)

plates (Tryptose blood in a modified Schaeffer’s Growth was initiated by culture reached 200 Klett inoculum was transferred

RNA

POLYXERASE

,4ND

105

SPORULATION

to fresh medium. This culture was allowed to grow to 200 Xlett units (late log phase) and used as a 5% (v/v) inoculum to initiate all experiments. The flask volume to medium volume ratio was always 10: 1 or greater. Cultures were vigorously aerated. (d) Mutagenesis WR746 spores were mutagenized &am & Waites (1968).

with

ethane

methyl

as described

sulfonate

(e) Selection of temperature sensitive

protease

by Mandel.

mutants

Mutagenized spores were plated onto 1% (w/v) skim milk-Tryptose plates a,t 47% Colonies which exhibited reduced zones of hydrolysis were replicated onto skim milkTryptose plates at 30 and at 47°C. A wild-type control also was spotted on each replica plate. Only those colonies giving reduced clearing at 47°C and normal clearing at 30°C were characterized further. (f) Estimation

ofreversionfrequency

Approximately 1 x lOa mutant cells were inoculated onto a Tryptose plate at 47°C. After 3 days incubation the small opaque colonies were replicated onto Tryptose plates at 30°C. Isolated single colonies were characterized further as to frequency of sporulation and hydrolysis characteristics at 30 and at 47°C. Direct selection for protease plus revertants was accomplished by plating 1 x lo* mutant cells onto a 2% (w/v) bovine serum albumin-Tryptose plate at 50°C. After 2 days incubation those colonies surrounded by large zones of clearing were streaked on Tryptose plates at 30°C. Isolated single colonies were further characterized as to their frequency of sporulation at 30 a,nd at 47°C.

(g) Estimation

of sporulation

frequency

The percentage of refractile bodies was quantitated as described & Doi, 1971). Heat resistance was measured by incubating cells The heat-resistant fraction was determined by comparing colony plates before and after heat treatment. (11) Antibiotic

activity

The antibiotic activity of cells grown on Tryptose mined by the method of Spizizen (1965). (i) Protease

previously (Leighton at 80°C for 29 min. counts on Tryptose

plates

at 30 and at 47°C was deter-

assay

low3 M-CaCl,, Commercial azocasein was dissolved as a 5% (w/v) solution in 0.2 x-Tris, pH 7.2. The solution was stored at -20°C. A l-ml reaction mixture contained: O-2 ml of azocasein solution, 0.2 ml of 1 M-Tris (pH 7*2), 0.1 ml of 10m2 M-f&&, an appropriate amount of protease solution and water to make 1 ml. The reaction was terminated by the addition of 1 ml of 10% (w/v) trichloroacetio acid. The reaction tubes were cooled at 0°C for 15 min and centrifuged to remove the precipitated protein. The supernatant fraction was carefully withdrawn and filtered through a 0.45 pm (11 mm) Millipore filter. A O.&ml amount of the filtrate was mixed with 0.2 ml of 1.8 N-NaOH and the absorbancy resd at 420 nm. Units of proteolytic activity are expressed as mg of azocasein hydrolyzed per hour. Reaction mixtures were incubated at 37°C except where otherwise indicated. The amount of proteolytic activity due to serine protease was determined by a difference assay. Two appropriate portions of protease solution were incubated at 30°C for 2 12. At the start of the incubation one of the portions was made 10m3 M with respect to phenyl methane sulfonyl fluoride. The amount of proteolytic activity inhibited by this compound was assumed to be due to serine protease. An identical procedure was used to estimate metal protease activit,y, except that the inhibitor was 10e3 M-o-phenanthroline.

Protein concentration bovine serum albumin

( j) Protein determination was estimated by the procedure as a standard.

of Lowry

et al. (1951) utilizing

106

.iZT AL.

T. J. LEIGHTON (k) Thermal denatzcration

conditions

All thermal denaturation experiments were done at 65% Culture supernatant fractions were from WB746, WB746 ts-5 and WB746 ts-5,rev-1 cells grown in Schaeffer’s medium for 8 h at 30°C. Cells were removed from the culture medium by centrifugation at 10,000 g for 10 min. The supernatant fractions were stored at - 20°C. Upon thawing, a precipitate formed; this precipitate was removed by centrifugation at 12,000 g for 10 min. Less was found in the precipitate. The supernatant than 5% of the total proteolytic activity fraction was filtered through a 0.45 pm Millipore filter. The filtrate was used without dilution for all denaturation experiments involving crude protease preparations. Pure serine protease was denatured in 0.2 M-Tris, 10y3 M-CalC12 (pH 7.2) at a protein concentration of 150 pg/ml.

(1) Pur$cation

of serine protease

WB746 and WB746 ts-5 cells were grown for 8 h at 30°C. The cells were removed by centrifugation at 10,000 g for 10 min. Four liters of culture supernatant fraction were adjusted to 0.65 saturation with respect to ammonium sulfate. The solution was allowed to stand overnight at 4°C. The precipitate was removed by centrifugation at 12,000 g for 15 min. The precipitated material was dissolved in 100 ml of 0.2 rvr-Tris, 10T3 M-&&l, (pH 8.0) and dialyzed exhaustively against the same buffer. The dialysate was brought to 0.65 saturation with respect to ammonium sulfate and allowed to stand at OV for at least 1 h. The precipitate was collected by oentrifugation at 15,000 g for 15 min. The precipitated material was dissolved in 10 ml of 0.01 iv-Tris, 1O-3 RI-CaCl, (pH 8.0) and exhaustively dialyzed against the same buffer. The dialysate was applied to a 2.0 cmx 40 cm DEAE-Sephadex (A50) column, previously equilibrated with 0.01 M-Tris, 10m3 MCaCI, (pH 8.0) and eluted with a 0.00 M to 0.15 M linear NaCl gradient,. The proteolytio activity eluting at approximately 0.03 ~-N&cl was pooled and brought to 0.75 saturation with respect to ammonium sulfate. The solution was allowed to stand overnight at 0°C. The precipitate was collected by centrifugation at 15,000 g for 30 min. The precipitated material was dissolved in 0.01 M-Tris-maleate, 10e3 M-C&l2 (pH 6.0) and exhaustively dialyzed against the same buffer. The dialysate was applied to a 2.0 cm x 40 cm CMcellulose-Sephadex (C50) column previously equilibrated with 0.01 M-Tris-maleate, (pH 6.0) and eluted with a 0.00 M to 0.30 M linear NaCl gradient. The 10-s M-Cd& proteolytic activity eluting at approximately 0.20 M-NaCl was pooled and brought to 0.75 saturation with respect to ammonium sulfate. The solution was allowed to stand overnight at 0°C. The precipitate was collected by centri.fugation at 15,000 g for 30 min. The precipitated material was dissolved in a minimal vol. (approximately 0.5 ml) of 0.2 M-Tris, 10L3 M-CaCl,, 2 M-Nacl (pH 8.0) and applied to a 2.0 cm X 80 cm G75 Sephadex column previously equilibrated with the same buffer. The fractions which corresponded to the 28,000 mol. wt region and contained proteolytic activity were dialyzed against 0.2 M-Tris, 10e3 M-Cd& (pH 7.2) and stored at -20°C. (m) Polyacrylamide

gel electrophoresis

Polyacrylamide gels containing sodium dodecyl sulfate were prepared according to the procedure of Laemmli (1970). Polyacrylamide gels containing urea were prepared according to the procedure of Bonner et al. (1968). All samples were exhaustively dialyzed against the appropriate application buffer. (n) RNA

polymerase

assay

RNA polymerase activity was assayed by a modification of the procedure of Harford & Sueoka (1970). A 0.25-ml assay mixture contained: 125 nmol of Tris (pH 8.0), 0.1 nmol of dithiothreitol, 37.5 nmol of KCl, 2.5 nmol of MgCl,, 0.5 nmol of MnCl,, 0.25 nmol each of ATP, CTP, UTP and GTP, 2 PCi of [3H]UTP (spec. act. 2.5 Ci/mmol), either 10 pg of de DNA or 20 pg of poly(dA-dT), an appropriate amount of enzyme (less than 75 pg), and water to make 0.25 ml. The reaction was stopped by the addition of 20 vol. 5% trichloroacetio acid containing 0.01 M-sodium pyrophosphate. Millipore glass-fiber filters were soaked in 5% trichloroacetic acid containing 0.01 M-sodium pyrophosphate before use. One unit of RNA polymerase activity incorporated 1 nmol of UMP per h at 37°C.

RNA

POLYMERASE

AND

107

SPORULATION

(0) $e DNA preparation +e BNA

was prepared

(p) Expression

from

high-titer

lysates

of the ts-5 phenotype

by the procedure

of Zubay

et al. (1970).

in the presence of chloramphen~col

WI3746 ts-5 cells were grown in Schaeffer’s medium for 5 h at 47°C. Chloramphenicol was added to a final concentration of 100 pg/ml. The culture was incubated for 10 min and one-half of the culture shifted down to 30°C; the other half of the culture remained at 47°C. Both portions were incubated for an additional hour. The experiment was terminat,ed by pouring the cells over ice. The preparation of cell-free extracts and crude RKA polymerase from these cells was as decribed by Losick & Sonenshein (1969). (q) In vitro

stability

of the ts-5 gene product

Cell-free extracts (Leighton & Doi, 1971) were prepared from WB746 and WB746 es-5 cells grown in Schaeffer’s medium for 5 h at 47°C. The extracts were incubated at 47 or at 30% and O*l-ml samples assayed for serine protease activity as described in section (i) above. In the cases where pure WB746 or WB746 ts-5 serine proteases were present the enzyme was added at 0 time.

(I-) Inhibition

of extracellular

spordation

serine protease activity during time periods

WB746 and WB746 ts-5 cells were grown in Schaeffer’s medium for 5 h at 47’C. The culture medium was made lo-* M with respect to phenyl methane sulfonyl fluoride and incubation continued for 10 min. The cells were collected by centrifugation at 10,000 g for 10 min. The pellet was resuspended in 0.01 M-Tris*HCl, 0.05 M-Mg& (pR 8.0) and centrifuged at 10,000 g for 10 min. This washing step was repeated 3 additional times. SubsequenUy, the cells were sonically disrupted (Leighton 85 Doi, 1971) and serine protease activity determined.

(s) E&erase assay contUitions Esterolytic activity against benzoyltyrosine ethyl ester was determined spectrophoto. metrically as described by Millet (1970). One unit of esterase activity hydrolyzes 1 pmol of substrate per min at 37°C.

3. Results (a) Xelection of temperature

sensitive protease mutants

It is clear that the isolation of totally negative hydrolytic mutants is going to select for mutations in genes other than those directly responsible for casein degradation (i.e. regulatory genes, membrane genes, auxotrophic genes, sporulation genes, etc.). Hence, we have screened for mutants which exhibited significantly reduced casein hydrolysis at a non-permissive temperature (4’7°C). A subclass of these mutants was then selected which produced normal zones of hydrolysis at a permissive temperature (30°C). The majority of such mutants (90%) were concomitantly temperature sensitive for sporulation. These mutants occurred at a frequency of 1 in lo5 survivors of the mutagenesis procedure. The remaining 10% of this subclass were oligosporogenous at 47°C. We have observed that mutants with slightly reduced zones of clearing at 47°C were not always asporogenous. One representative of the major class of temperaEure sensitive hydrolytic mutants, WB746 ts-5, was chosen for further study. (b) The ts-5 phenotype is due to a single-site mutation Revertants with normal hydrolytic properties at 47°C cannot be directly selected on skim milk-Tryptose plates since WB746 ts-5 is not a totally negative mutant at

108

T. J. LEIGHTON

ET’

AL.

this temperature. However, revertants with wild-type sporulation frequencies can be selected easily at 47°C. Mutants blocked early in sporulation lyse on Tryptose plates after prolonged incubation. This effect can be exploited to select sporogeneous revertants since these cells will form small colonies (overgrow) after incubation for several days. Hence, if lo8 WB746 ts-5 cells are plated on a Tryptose plate at 47°C and incubated for three days, spontaneous revertants appear at a frequency of 1 in lo7 cells. The revertants such as WB746 ts-5,rev-I, sporulated at high frequency and had normal hydrolytic properties at 47°C (Table 1). The serine protease thermal stability of one typical revertant, WB746 ts-5,rew1, will be discussed later. Protease plus revertants can be directly selected for on 2% bovine serum albumin-Tryptose plates at 50°C. Under these conditions WB746 h-5 cells appear to be protease negative. Protease plus revertants appeared at a frequency of 1 in 10’ cells. In the majority of such revertants (95%) sporulation became temperature independent. The properties of one such revertant, WB746 ts-5,rev-10, are described in Table 1. The 5% of the revertants which became protease plus, but which were still temperature sensitive for sporulation, represent reversion events which increased the extracellular levels of the temperature sensitive enzyme. We have examined the thermal denaturation properties of several of these revertants. The serine protease produced by these strains had the same thermal stability as the h-5 enzyme (data not shown). We have further attempted to establish that the lesion causing temperature sensitive casein hydrolysis is the same locus responsible for temperature sensitive sporulation. Since WB746 ts-5 was prototrophic, as was the parent strain WB746, we have prepared bacteriophage PBS1 transducing lysates from WB746 ts-5 and used these lysates to transduce various auxotrophic strains of B. subtilis to prototrophy (Van Alstyne & Simon, 1971). In the case of strains SBIZO, an aromatic amino acid auxotroph, and BRl6, a lysine auxotroph, we found that a significant portion of the transductants were concomitantly temperature sensitive for casein hydrolysis. In addition, in every case (46/46) where we found a transductant which was temperature sensitive for casein hydrolysis this strain also was temperature sensitive for sporulation. Recombination frequencies from these two-factor crosses indicated that the ts-5 mutation was 30% cotransduced with aroD and 1~8-1. This would place the ts-5 lesion close to a large cluster of sporulation mutants (Young & Wilson, 1972) and a streptomycin resistance marker, strC2 (Staal & Hoch, 1972). We have prepared transducing lysates from strC2 and used these preparations to transduce WB746 ts-5 to streptomycin resistance. We observed that 85% of the transductants which received the strC2 marker became temperature independent for sporulation. These two types of transduction experiments give complementary results and establish that the h-5 marker behaves as a single-site mutation. We have made no attempt to order the h-5 marker wit.h respect to neighboring markers such as strC2 or spoAl2. Since we are not sure that either the strC2 or the spoAl2 strains are completely isogenic with respect to WB746, any fine-structure mapping will have to await the construction of appropriate isogenic strains of WB746 carrying these markers. Also, we would like to have additional tightly linked selective markers so that the more accurate method of transformational mapping could be used. Even though the strC2 and spoA12 mutations are tightly linked by cotransduction these markers cannot be linked by transformation (Staal & Hoch, 1972).

1

45 + -I0.95 + ++

Generation time Casein hydrolysis Bovine serum albumin hydrolysis Fraction of resistant cells Antibiotic activity Esterase activity -,+,-

: 0.90

T 0.92 +++

45

45

WB746 ts-5

j: 0.90 __ t’+

45

WB746 ts-5, rev-10

-

-_

spoA12

28 -i- + ++ 0.80 + -t

WB746

Temperature

28 zt 10-T i-

WB746 ts-5

WB746 ts-5,rev-10;

i-

E 0,70

28

WB746 ts-5, rev-l

47 or 50°c

spoA12

:I 0.75 i +

28

WB746 h-5, rev-10

-

-

spoA12

Experimental conditions are described in the text. All characteristias were measured at 47°C except the albumin hydrolysis, which was measured at 50°C. Plates usod to determine 813012 chsract,eristics were supplemented with 20 log tryptophan/ml. (f +) E&erase activity, 1027 units/ml; (+) osterase activity, 604 units/ml; standard deviation among the various strains is &loo/, of these values.

WB746

Characteristic

3o”c WB746 h-5, rev-l

Characteristics of WB746; WB746 ts-5; WB746 ts-5, rev-l;

TABLE

110

T. J. LEIGHTON

E2’

AL.

(c) Thermal denaturation of WBY46, WB746 ts-5 and WBY46 ts-5,rev-I serine proteases Using the purification procedure for serine protease activity described above we recovered 55o/o of the activity originally present in the culture supernatant fraction. This represents 12 mg of pure serine protease from four liters of starting material. Approximately 90% of the culture supernatant fraction activity was recovered after the ammonium sulfate step ; 80% of the original activity was recovered in the pooled DEAE-Sephadex fractions; 65% of the original activity was recovered in the pooled CM-cellulose-Sephadex fractions ; and 55% of the original activity was recovered in the pooled G75 fractions. Figure 1 illustrates that in crude or pure preparations ts-5 serine protease was inactivated significantly faster than wild-type protease at 65°C. In addition, the serine protease from a single-step revertant was much closer to wild-type protease in its thermal stability. (d) Electrophoretic mobility of pure WBY46 and WBY46 ts-5 se&e proteases When pure preparations of WB746 and WB746 ts-5 serine proteases were electrophoresed in 10 cm pH 4.5 urea geIs for eight hours the ts-5 protease migrated significantly faster. Two bands were evident in the co-electrophoresis experiment shown in Figure 2. We should point out that the densitometer was adjusted to zero on the unstained areas adjacent to the protease bands in order to aid integration and visual presentation. On prolonged storage of the protease we have observed the appearance of several faster moving bands not present in the original preparation. We assume this is due to autodigestion known to occur in subtilisin preparations (Ottesen &

(a)

Ttme (min)

(b)

FIG. 1. Thermal inactivation of WB746, WB746 ts-5 and WE5746 ts-5,rev-1 serine proteases at 66°C. Experimental conditions are described in the text. (a) Den&w&ion of swine protease in crude-culture supernatant fractions. -e--e---, WB746 enzyme; --m--m--, WB746 ts-5 enzyme; -A-A--, WB746 ts-5,rev-l enzyme. (b) DenaturaWB746; --m--I--, WB746 ts-5. tion of pure se&e protease. -.-.-,

RNA

POLYMERASE

AND

SPORIJLATION WB

74,

I 5

6 Migration

2 (cm)

5 (b)

(a)

FIG. 2. Electrophoretic mobility of pure WB746 and WB746 ts-5 serine proteases in pN 4.5 7.5% polyacrylamide urea gels. Experimental conditions are described in the text. (a) Co-electrophoresis of pure WB746 and WB746 h-5 serine protease. Gels (10 cm) were run for 8 h at 5 mA/gel. No bands are seen on these gels other than the doublet at the position indicated. Identity of the two bands was confirmed by running each enzyme separately in duplicate gels. (b) Co-electrophoresis of phenyl methane sulfonyl fluoride-treated pure WB746 and WB746 ts-5 serine protease. Conditions were identical to those described for (a). Approximately 2 pg of each enzyme was applied to the gels.

Svendsen, 1970). In a few preparations we have observed a minor contaminant (274 of the total protein applied to the gel), which runs at approximately 7.6 cm in this system. This protein was absent from preparations having full enzyme activity. The basic metal protease runs in this position approximately. Since we have observed that all the protease activity in pure preparations was fluoride inhibitable, this band

!IIIJIJIII/ 0 01 0.2 03

04 05 06 07 08 09

Relatw

mobility

FIG. 3. Molecular weight estimation of WB746 and WB746 ts-5 serine proteases in 7.5% polyacrylamide, 0.1% sodium dodecyl sulfate gels. The gels w ere run as described by Laemmli (1970) Each protease was co-electrophoresed with the following standards: trypsin (23,000), subtilisin (28,500), aldolase (40,000), serum albumin (68,000) and phosphorylase A (94,000). (m) Indicates the relat’ive mobility of both enzymes in this system.

112

T. J. LEIGHTON

ET

Al;.

may represent inactive metal protease or alternatively a small amount of autodigestion. The fact that the bands seen on these gels were due to serine protease rather than a contaminant is demonstrated in Figure 2(b). In this co-electrophoresis experiment both proteases were preincubated with lo- 3 M-phenyl methane sulfonyl fluoride for two hours at 30°C before dialysis against electrophoresis buffer. The mobility of both peaks was significantly retarded after reaction with the inhibitor. Wild type and ts-5 proteases, if not inhibitor treated, must be dialyzed separately and mixed immediately before electrophoresis, since during dialysis the wild-type protease will cleave the ts-5 enzyme into several faster-moving bands. Co-electrophoresis of heat and sodium dodecyl sulfate denatured proteases from wild type and ts-5 cells suggests a minimum molecular weight of 28,OOO,f500 for both enzymes (Fig. 3). There was no noticeable mobility difference between the two enzymes when co-eleotrophoresed in this gel system. Since the two proteases differ only in charge, and not size, it is possible that the ts-5 mutation is a single-site missense lesion. (e) SpeciJic activity and temperature dependence of WB746, WB746 ts-5 serine proteases Table 2 lists the average specific activity of six different preparations of WB746 and WB746 ts-5 serine proteases. These enzymes differed significantly in their specific activities against azocasein at 30°C. Furthermore, WB746 specific activity increased 400% for an in vitro shift from 30°C to 47°C. WB746 ts-5 serine protease activity increased only 300% over a similar increment. Approximately 50 to 60% (variation for 6 preparations) of the total serine protease activity in crude supernatant fractions was recovered as pure enzyme. TABLE

2

Specific activity of pure WB746 and WB746 ts-5 serine proteases Source of enzyme

WB746 WB746 b-5

Specific activityt at 30°C

Specific activity?

750 600

Experimental conditions are described in the text. Standard t Specific activity is in units/mg protein/h.

at 47°C 3000 1800 deviation

is 17%.

(f) Serine and metal protease activity levels during growth at 30°C and 4r”C Figures 4 and 5 depict the time-course of protease accumulation in wild type and ts-5 cells at 30°C and at 47°C. It is evident that the intracellular and extracellular accumulation profiles were temporarily identical. Protease accumulation was initiated at the end of the exponential growth period and increased at a linear rate until shortly before refractile bodies appeared. On a units/ml basis approximately I to 2% of the total proteolytic activity was found inside the cell. This is a very significant amount of activity when one considers the ratio of intracellular volume to culture medium volume, In addition, in the case of ts-5 cells, the decrease in serine protease

RNA

POLYMERASE

AND

SPORULATION

I :n

activity seen at 47°C was identical whether intracellular or extracellular levels were compared. At 47°C WE746 ts-5 cells produced normal levels of metal protease (Fig. 5(b)) and serine esterase (Table 1). Hence, the only extracellular or intracellu1a.r in h-5 cells was a serine enzyme. It, can be argued protease which was underproduced

!

0.6 i O-4 i I

0.2

I I I

0

I

I I

I I

I23456789101112

I I

I

Illil~llllll

I

0 I23456769101112 Time (hl

4. Extracellular accumulation of serine and metal protease activity in WB746 and WB746 ts-5. Experimental conditions are as described in the text. (a) Serine protease accumulation at 47°C. -e-a--, WB746; --m--m--, WB746 ts-5. (b) Metal protease accumulation at 47°C. -a--a---, WB746; --m--H--, WB746 ts-5. (c) Serine protease accumulation at 30°C. (0) WB746; ( n ) WB746 h-5. (d) Metal protease accumui&ion at 30°C. (e) WB746; (m) WB746 ts-5. FIG.

that the intracellular activity we observe is actually extjracellular during the sonic disruption process necessary to prepare cell-free is not, the case is demonstrated in Table 3. When a concentration to inhibit greater than 950/ of the extracellular serine protease 8

enzyme solubilized extracts. That this of fluoride sufficient activity was added

114

T. J. LEIGHTON

ET

AL.

to WB746 and WB746 k-5 cells grown at 47”C, and precautions were taken to remove the inhibitor prior to cell breakage, intracellular protease levels similar to untreated cultures were obtained. Hence, although all the extracellular serine protease had been inactivated there still were normal amounts of intracellular serine enzyme.

c

[b)

I23456789101112 Time (h)

5. Intracellular accumulation of serine and metal protease activity in WB746 and WB746 ts-5. Experimental conditions are described in the text. Figure designations and symbols are identical to those in Fig. 4. FIG.

Also, the ratio of serine enzyme activity to metal enzyme was unchanged. All protease assays were run at 3O”C, and since the b-5 enzyme had an abnormal temperature dependence, the activity values for 47°C b-5 cells were an over-estimation of the actual enzyme activity at this temperature. These experiments estimate the maximum potential proteolytic activity present at 47°C.

RNA

POLYMERASE

AND TABLE

IP.5

SPORULATION

3

Ejfect of phenyl methune suljonyl jluoride PretreutmeTaton intracellular protease levels Source of cells

WE-746 WB746 + inhibitor WB746 b-5 WB746 h-5 + inhibitor

Specific activity? of serine enzyme

Specific activity? of metal enzyme

0.48 0.40 0.12 0.10

Experimental conditions are described in the text. Standard T Speoific activity is in unitsjmg protein/h at 30°C.

0.12 0.10 0.17 0.15 deviation

is & 10%.

(g) In vitro stability of WB746 and WB746 t,s-5 serine proteases Table 4 illustrates that pure wild type and ts-5 extracellular enzymes prepared from cells grown at 30°C were completely stable to incubation at 30°C and 47°C. This result’ does not offer any explanation as to why we find reduced levels of serine protease in ts-5 cells grown at 47°C. Subsequent experiments revealed that the k-5 serine protease underwent an abnormal conformational change between 30°C and 47°C which made the enzyme more labile to degradation or inactivation by some other system present in wild type and ts-5 cells. Hence, in crude cell-free extracts ts-5 serine protease activity was stable over long periods at 30°C but was rapidly destroyed. at 47°C. Evidently in vivo serine protease synthesis was somewhat faster than degradation since we could assay serine protease activity in k-5 cells which was subsequently destroyed in vitro. We present further evidence to support this point in the following section. (h) Expression of the ts-5 gene product in the absence of protein synthesis The most reasonable explanation for the result that ts-5 cells grown at 47°C had low levels of serine protease which could be assayed as active against azocasein, but which did not result in any detectable modification of RNA polymerase, was that the abnormal conformational change which this protease underwent between 30°C and 47°C (which made the enzyme more labile in vivo) resulted in a loss of activity against its natural substrate, RNA polymerase. Hence, the ts-5 mutation is in essence a temperature sensitive substrate specificity lesion. This hypothesis can be tested easily by shifting down post-exponential ts-5 cells grown at 47°C under conditions (Leighton & Doi, 1971) which totally inhibit protein synthesis. The results of such an experiment are presented in Table 5. Post-exponential WB746 ts-5 cells which remained at 47°C after exposure to chloramphenicol retained the vegetative template specificity. However, k-5 cells which were shifted down to 30°C after exposure to chloramphenicol, rapidly changed to sporulation-type template specificity. Since there ha.d been no de novo protein synthesis after the temperature shift, the ts-5 protease which was “inactive” against RNA polymerase at 47°C had become active at 30°C. We would presume that this indicates that the k-5 serine protease had changed from an inactive to an active conformation which was capable of cleaving the RNA polymerase ,&subunit. In addition, these results establish that the k-5

116

T. J. LEIGHTON

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TABLE 4 In vitro

stability

of WB’i’46 and WBY46 ts-5 serine proteases

Serine enzyme preparation

0 time

Serine protease specific activity? 80 min incubation 320 min incubation at 30°C at 47°C 733 601 0.41

730 599 0.40

728 597 0.45

Pure WB746 Pure WB746 b-5 WB746 cell-free extract WB746 k-5 cell-free extract WB746 cell-free extract f 0.5 unit pure WB746 protease/mg protein WB746 cell-free extract + 0.5 unit pure h-5 protease/mg protein WB746 ts-5 cell-free extract + 0.5 unit pure WB746 ts-5 protease/ mg protein WB746 ts-5 cell-free extract + 0.5 unit pure WB746 protease/mg protein

0.12

0.01

0.10

0.95

0.92

0.94

0.97

0.42

0.93

0.62

o-04

0.58

0.62

0.48

0.60

Experimental procedures are described in the text. When pure serine protease was added to crude extracts this was done at 0 time. Standard deviation for pure enzyme assays is &7% and for crude enzyme assays & 10%. ? Speoifio activity is in u.nits/mg protein/h at 30°C.

mutation is not a regulatory lesion which requires translation to be expressed. These data are in complete agreement with the previous experiments, suggesting that the h-5 phenotype is due to a mutation in the structural gene coding for the serine protease. TABLE 5 The expression

of the ts-5 phenotype

in the presence of chloramphenicol Specific activityf Poly(dA-dT) de

Source of enzyme 47°C post-logarithmic chloramphenicol/ml 47°C post-logarithmic chloramphenicol/ml

WB746 ts-5 cells and held at 47°C WB746 ts-5 cells and shifted down

+ 100 pg for 1 h + 100 pg to 30°C for 1 h

14.6

4.0

1.0

3.2

Experimental procedures are described in the text. WB746 ts-5 cells were grown for 5 h at 47’C before initiating the experiment. Standard deviation is f 10%. t Specific activity is in units/mg protein at 37°C for ammonium sulfate-stage RNA polymerase.

4. Discussion It is clear from our previously published data that the absence of a functional serine protease results in cells which are blocked biochemically and physiologically at or near stage 0 in the sporulation sequence (Leight,on et al., 1972). Electron microscopic examination of ts-5 cells grown at the non-permissive temperature (Santo et al., 19721 suggests that these cells also are blocked morphologically at st’age 0.

RNA

POLYMERASE

AND

SPORULATION

11%

Numerous criteria strongly suggest that the ts-5 lesion is a single-site mutation in the structural gene coding for a basic serine protease. This protein is found as an extracellular enzyme. Obviously the enzyme must be synthesized intracellularly, and our data would argue that some form of the ts-5 gene product must persist in the intracellular environment. The fact that intracellular and extracellular serine protease levels are reduced to the same extent in k-5 cells grown at 47°C would argue for a relation between these activities. Furthermore, since the is-5 enzyme is stable at 47°C in the ext,racellular environment, and unstable at 47°C in the intracellular environment, the only way in which extracellular enzyme levels could be reduced is by degradation or inactivation of the enzyme intracellularly. Prestidge et al. (1971) also have noted a direct correlation between the extracellular and intracellular levels of proteases during sporulation in B. subtilis. Whether most of the serine protease &ivity found intracellular is the ts-5 gene product remains an open question, since the observed reduction in intracellular serine protease levels could be an indirect resuh of the ts-5 mutation. Our method of estimating intracellular serine proteaso activity depends upon using phenyl methane sulfonyl fluoride to inactivate the ext,raeellular serine protease, and subsequent removal of the inhibitor prior to cell breakage. It is clear that under our conditions there is no significant inhibition of int.racellular serine protease activity by this fluoride. However, it should be pointed out that exposure of Escherichia coli cells to much higher concentrations of fluoride for longer periods of time, does appear to inhibit some intracellular proteases (Goldberg, I971a). Recently Shoer & Rappaport (1972) have described an extracellular serine protease mutant of B. subtilis which can sporulate. This acidic protease has a very different amino acid composition (Shoer & Rappaport, 1972) and electrophoretic mobility (Rappaport et al., 1965; Riggsby & Rappaport, 1965) from the basic serine enzyme studied in this investigation (Boyer & Carlton, 1968; Millet, 1970; Ottesen & Svendsen, 1970). It is possible that this enzyme is not required for dporulation and may be related to the acidic serine e&erase described by Millet (1970) or the acidic serine protease described by Boyer & Carlton (1968). Th.e reasons for the temperature sensitive phenotype of ts-5 cells are rather complex. Pure ts-5 serine protease produced at the permissive temperature can readily be distinguished from wild-type protease on the basis of specific activity, thermal stability at 65”C, mobility in pH 4+5urea gels and temperature dependence. However, pure ts-5 protease isolated from culture supernatants is completely stable at 47°C: as is the wild-type protease. Hence, denaturation per se, is not an adequate explanation for the reduced intracellular and extracellular protease levels found at 47°C. However, in crude cell-free extracts k-5 serine protease activity is rapidly destroyed at, 47°C. At 30°C the serine protease activity is stable over long periods. It appears that the abnormal conformational change which the ts-5 enzyme undergoes between 30°C and 47°C makes the enzyme more labile to an inactivation process in these cells. The results of Goldberg (1971b,1972), which indicate missense proteins can be selectively degraded in viva, may be very pertinent to this observation, A possible explanation for the reduced protease levels in ts-5 cells grown at 47°C is that the mutant protease is in a conformation which is recognized as an a,bnorma,l protein and degraded by turnover systems known to be operative in these cells. Several types of evidence at also suggest that when the ts-5 serine protease is in the abnormal conformation 47°C it cannot recognize its natural substrate, RNA polymerase. Since the B. subtilis basic serine protease is closely related to subtilisin (Hageman

IlS

T. J. LEIGHTON

ET

AL.

& Carlton, 1970), this mechanism for temperature sensitivity is most appealing. Subtilisin is known to contain no cysteine or eystine, and to have a very low level of a-helical content (Ottesen & Svendsen, 1970). These structural simplicities would make the isolation of a temperature sensitive mutation which makes the molecule more labile to spontaneous denaturation at physiological temperatures extremely unlikely. Subtilisin is known to produce very selective proteolytic cleavages of native proteins (Ottesen & Svendsen, 1970). The findings that subtilisin and an intracellular protease of B. subtilis are both capable of selectively cleaving E. coli DNA polymerase (Brutlag et al., 1969; Setlow et al., 1972; Setlow & Korn’berg, 1972) are consistent with our data. Temperature sensitive mutants are a well documented means of isolating lesions affecting a structural gene of interest. Such mutants provide a critical biological control which establishes whether the gene product has any importance under normal physiological conditions. We have considered in great detail an argument not often recognized in studies of this type which would contend that the temperature sensitive mutation affects some other molecule which interacts in some way with the gene product assumed to be temperature sensitive. We believe that even given numerous ad hoc assumptions this type of argument cannot adequately account for our data, particularly the experiments in which the ts-5 mutation is correctly expressed in the absence of protein synthesis, after a shift to the permissive condition. Given that the h-5 mutation resides in the serine protease structural gene, it is evident that this protein is intimately associated with the conversion of DNAdependent RNA polymerase ,E subunit from the vegetative to the sporulation form. This association could be direct or indirect. Experiments done by P. K. Freese (Leighton et aZ., 1972; P. K. Freese & R. H. Doi, unpublished results) indicate that pure serine protease is capable of modifying vegetative RNA polymerase ,6 subunit, by a limit cleavage, to the 110,000 mol. wt form found in early sporulating cells (Losick et al., 1970). Concomitant with this molecular weight reduction is a loss of the ability to transcribe $e DNA, a template only used by vegetative RNA polymerase (Losick & Sonenshein, 1969). The in. vitro modified enzyme continues to transcribe poly(dA-dT), as does RNA polymerase isolated from sporulating cells. It would thus appear that the serine protease affected by the ts-5 mutation is directly responsible for the /l subunit molecular weight change characteristio of the transition from vegetative to sporulation time periods. This result is certainly not without precedent as Sadoff et al. (1970) have shown that a similar serine protease is capable of converting vegetative fructose 1,6-diphosphate aldolase to a protein indistinguishable from its spore counterpart. An identical type of experiment can be done under conditions which more closely approximate those in sporulating cells. WB746 ts-5 cells can be grown at 47°C for several hours post-exponentially with no change in the vegetative-type template activity. However, if chloramphenicol is added to these cells in sufficient concentration to totally inhibit protein synthesis (Leighton & Doi, 1971), and the culture is shifted down to 3O”C, there is a rapid conversion of RNA polymerase from the vegetative to sporulation form. A satellite culture treated in an identical manner, and held at 47°C retains the vegetative template specificity. Since the only molecule which has changed state during the shift down is the serine protease, this should give a result similar to the in vitro modification. In fact, the two experiments give identical changes in template activity. We should point out there is no absolut,e increase in

RNAPOLYMERASEAND

SPORULATION

119

poly(dA-dT) specific activity in either experiment even after the enzyme has become totally inactive on #e DNA. A careful study of the time-course of poly(dA-dT) specific activity changes (Leighton, unpublished data) reveals that the increased a,ctivity on this template occurs in vivo after RNA polymesase has been completely converted to the 110,000 molecular weight species. Hence, all of these results establish that the in vitro and in situ systems give identical results with those found in viva. The chloramphenieol experiments also suggest that the ts-5 protease is in a conformation at 47°C which cannot cleave RNA polymerase although it can hydrolyze casein. Upon shift down to 30°C the enzyme returns to a normal conformation modifies RNA polymerase. Hence the ts-5 mutant is a temperature sensitive spec city .mutation in the serine protease. This explains why we can detect significant amounts of serine protease activity in h-5 cells grown at 47”C, but observe no change 6n RNA polymerase template specificity. On the basis of these results we predicted it should be possible to isolate an RNA polymerase conformation mutant which would suppress the ts-5 mutation at 47°C. This double mutant should allow sporulaLion to proceed past stage 0. Such a mutant has been found and its properties will be discussed in detail later (Leighton, unpublished data). The reeent studies of Brevet & Sonenshein (1972) are complementary to our interpretation of the ts-5 phenotype. They have found that eight stage 0 sporulation mutants of B. subtilis, which are lacking the basic serine protease, fail to undergo the sporulation-specific change in RNA polymerase template specificity. Sporulation -mutants which are blocked at later stages do exhibit the characteristic change in template specificity. Furthermore, one stage 0 mutant, spoO-4u, which contains near wild-type levels of serine protease, does exhibit a significant change in RNA polymerase template specificity. Our results would suggest that in this mutant other sporuiation-specific processes must be affected which stop sporulation at stage 0. In contrast to our results Millet, et al. (1972) claim that an intracellular protease from B. megaterium is capable of modifying B. subtilis RNA polymerase to the sporulation form, and that the basic serine protease non-selectively degrades RNA polymerase. Several features of this report deserve comment. The non-specific action of the basic serine protease is inconsistent with our biochemical and genetic data, as well as the data of Sadoff et al. (1970), B&lag et ab. (1969), Setlow & Kornberg (1972), and Setlow et al. (1972). Since Millet et al. (1972) have not shown any analytical purity data for their serine protease preparation, it is possible that this material could be contaminated with other extracellular proteolytic enzymes. Also, there are no data included as to the effect of muoh lower amounts of basic serine protease on RNA polymerase structure. Furthermore, these authors demonstrate t,hat subsequent to modification a new RNA polymerase subunit appears in the 130,000 molecular weight region, and that this modified enzyme has decreased activity on +e DNA and greatly increased activity on poly(dA-dT). Maia et al. (1971) have reporte that this form of RNA polymerase is found in B. subtilis spores. These results are totally inconsistent with all the known biochemical (Losick & Sonenshein, 1969; Brevet $ Sonenshein, 1972; Losick et al., 1970; Losick, 1972; Leighton et al., 1972) and genetic (Leighton & Doi, Abst. Ann. Meetings Am. Xoc. Microbid., 1972, p, and Leighton & Doi, unpublished data) facts associated with the state of R polymerase in sporulating cells. The modified form of RNA polymerase found in spowlating cells contains a 110,000 molecular weight /3 subunit (Losick eCd., 1970 ; Leighton et a%., 1972) rather than the 130,000 molecular weight /3 subunit reported

120

T. J. LEIGHTON

ET

AL.

by Maia et al. (1971). Losick et al. (1970) have shown t’hat partially purified RNA polymerase does contain a contaminating protein of 120,000 molecular weight. The relation of this contaminating protein to the subunit reported by Maia et al. (1971) is not clear. Furthermore, it is known that the unmodified form of RNA polymerase is present in B. subtilis spores (Losick, 1972; Leight,on & Hoi, Abst. Ann. Ne&ngs Am. SOC. Microbid., 1972, p. 63). Also, the serine protease which is capable of converting vegetative fructose 1,6-diphosphate adolase to the sporulation form is absent from spores (H. L. Sadoff, personal communication). The B. megaterium serine protease used by Millet et al. (1972) was purified from spores. In addition, from the data presented here and previously (Leighton et al., 1972), and from the results of Brevet & Sonenshein (1972), it is evident that subsequent to the modification of RNA polymerase there is no increase in the specific activity of this enzyme on poly(dA-dT). This increase occurs after the p subunit modification (Leighton, unpublished data; Losick & Sonenshein, 1969), and near the time when RNA polymerase is returning to the vegetative form. It is apparent that WB746 ts-5 is not similar to any previously isolated mutant which blocks sporulation at stage 0. All other stage 0 mutants fail to produce several of the extracellular enzymes found in B. subtilis culture supernatants (Michel & Millet, 1970; Guespin-Michel, 1971; Ito & Spizizen, 1972). WB746 ts-5 cells only underproduce serine protease activity when grown at 47°C. The ts-5 mutant can be easily distinguished from the spoA12 strain by the ability of ts-5 cells to hydrolyze casein at 47”C, spoAl2 cells give no zone of clearing on these plat,es. Although the ts-5 mutation maps in a region of the B. subtilis chromosome containing spoA12 a,nd related mutations, the ts-5 lesion must occupy a distinctly different genetic site. A test for the genetic identity of the ts-5 and spoA strains, by the recombination index method (Hoch, 1971), suggests these two markers are not in the same gene. (J. A. Hoch, personal communication.) Also, the fact that nearly every serine protease plus revertant is also sporulation temperature independent is not the type of reversion pattern seen with the other pleiotropic stage 0 mutants (Guespin-Michel, 1971). The complexity of the regulatory events controlling the early stages of sporulation is well demonstrated by our results. Mutations controlling the expression of the serine protease gene, which by itself has pleiotropic effects, map at several sites on t.he chromosome (Guespin-Michel, 1971; Ito & Spizizen, 1972; Brevet & Sonenshein, 1972), and exhibit complex partial reversion patterns (Guespin-Michel, 1971). It seems reasonable to assume that all of these mutations affect the expression of many genes necessary to complete the initial stages of sporulation. Some of these effects must be mediated by diffusible gene products as the spoOA mutants are dominant to the wild-type allele (Kermazyn et al., 1972). A superficially perplexing question is why the serine protease also is found as an extracellular enzyme. One could argue that the primary extracellular function is nutritional. The serine protease could act in conjunction with the other extracellular enzymes to provide a source of amino acids to cells which are attempting to survive under starvation conditions. An additional intriguing possibility is suggested from a preliminary analysis of the RNA polymerase mutant which suppresses the ts-5 phenotype at 47°C (Leighton, unpublished data). The double mutant cells proceed normally through the sporulation sequence at 47°C until the spore coat. deposition step is reached. At this stage, development becomes abnormal resulting in many spores being produced with extreme structural defects. These dat,a, in conjunction

RN-4 POLYMERASE

AND

SPORULaTION

121

with the fact that this serine protease cannot be detected in spores (II. L. Sadoff, personal communication), and the fact that RNA polymerase is in the vegetative subunit form at this stage (Losick, 1972), suggest that the serine protease may have a role in spore coat protein assembly. These results also would infer that all the active serine protease is in an extracellular environment during the final stages of spore production. Since coat deposition occurs in what is essentially an ext,racellular space between the mother cell and the forespore, this readily explains why the enzyme is found extracellularly and why a shift up of ts-5 cells late in sporulation, when RXA polymerase is not being modified, results in a stoppage of spore maturation. Experiments are in progess to clarify further the role of serine protease in spore coat morphogenesis. It should be noted that proteolytic cleavage of precursor proteins in bacteriophage T4 morphogenesis is a well established mechanism for assembly of a complex molecular structure (Laemmli, 1970). The modification of pre-existing vegetative proteins to new species which have altered specificity is an extremely appealing mechanism for molecular differentiation. This means of differentiation is very useful in terms of cell economy, especially when one molecule is capable of modifying several different proteins of developmental importance. Recent reports suggest that B. subtilis may not be the only organism to utilize such a system for the regulation of transcriptional specificity (Weaver et al., 1971). We are grateful to R. Losick for many stimulating discussions, and to J. Hoch for bacterial strains. This research was supported by funds from a General Research Services grant to the University of Massachusetts Medical School for one of us (T. J. L.), a National Science Foundation grant (GB-8405) and a United States Atomic Energy Commission grant (AT(O4-3)-34) to another of the authors (R. H. D.), and a National Research CoLmeil of Canada grant (NRC-A3686) to one of us (R. A. J. W.). REFERENCES Aronson, A. E., angelo, N. & Holt, S. C. (1971). J. Bact. 106, 1016. BaJassa, G. (1969). Molec. Gen. Genetics, 104, 73. Bernlohr, R. W. & Clark, V. (1971). J. Bact. 105, 276. Bonner, J., Chalkley, 6. R., Dahmus, M., Fambrough, D., Fugimura, F., Huang, R. C., Huberman, J., Jensen, R., Marushige, K., Ohlenbasch, H., Olivera, B. & Widholm, J. (1968). In Methods in Enzymology, ed. by L. Grossman & K. Moldave, p. 3, vol, 12. Academic Press, New York. Bayer, H. W. & Carlton, B. (1968). Arch. Biochem. Biophys. 128, 442. Brevet, J. & Sonenshein, A. L. (1972). J. Bact. 112, 1270. Brutlag, D., Atkinson, M. R., Setlow, P. & Kornberg, A. (1969). Biochem. Biophys. Res. Commulz. 37, 982. Goldberg, A. L. (1971a). Nature, 234, 51. Goldberg, A. L. (1971b). Proc. Nat. Acad. Ski., Wash. 68, 362. Goldberg, A. I.,. (1972). Proc. Nat. Aead. Sci., Wash. 69, 442. Guespin-Michel, J. P. (1971). J. Bact. 108, 241. Hageman, J. H. & Carlton, B. C. (1970). Arch. Biochem. Biophys. 139, 67. Harford, N. & Sueoka, N. (1970). J. Mol. Biol. 51, 267. Ho&, J. A. (1971). J. Bact. 105, 896. Hoch, J. A. & Spizizen, J. (1969). In #pores IV, ed. by L. L. Campbell, p. 112. Americnn Society for Microbiology. Ito, J. & Spizizen, J. (1972). In Spores V, ed. by H. 0. Halvorson, R. Hanson & L. L. Campbell, p. 107. American Society for Microbiology. Kermazyn, C., Anagnostopoulos, C. & Schaeffer, P. (1972). In spores TV, ed. by H. 0. Nalvorson, R. Hanson & L. L. Campbell, p. 126. American Society for Microbiology.

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Young, F. E. & Wilson, G. A. (1972). In Spores V, ed. by H. 0. Halvorson, R. Hanson & L. L. Campbell, p. 77. American Society for Microbiology. Zubay, G., Chambers, D. A. & Cheong, L. C. (1970). In 17he Lactose Operon, ed. by J. R. Backwith & D. Zipser, p. 375. Cold Spring Harbor Laboratories, New York.