Protein Degradation and Proteolytic Modification
Chapter
62
CHARLES G. MILLER
The hydrolysis of peptide bonds is a central metabolic activity in bacterial cells. Escherichia coli contains at least 40 enzymes that catalyze this reaction with representatives in every cellular compartment. These enzymes participate in a diverse array of processes, ranging from the catabolism of peptides to the specific inactivation of regulatory proteins.
The early studies that established the importance of protein degradation in E. coli were reviewed in the previous edition of this book (151). This work led to the concept that most E. coli proteins are stable in growing cells but that a large fraction of these proteins become susceptible to degradation when cells are starved. Growing cells are capable of protein degradation, however, since a variety of "abnormal" proteins (e.g., proteins containing amino acid analogs such as canavanine or protein fragments produced in cells carrying nonsense mutations) can be rapidly and specifically degraded during growth. Both of these degradative processes require metabolic energy.
For the most part, these concepts are still valid. There is clearly much more protein breakdown in growing cells than the early workers envisaged. Several proteins are extremely rapidly degraded during normal growth, and a significant fraction of newly synthesized peptide bonds are rapidly hydrolyzed. In most cases, these proteolytically unstable proteins act as regulators and their susceptibility to proteolysis plays a key role in this regulatory function. The list of proteins qualifying as abnormal has expanded substantially with the addition of cloned foreign proteins and fusion proteins. The requirement for energy is still a key feature of in vivo protein degradation, and great progress has been made in understanding its enzymatic basis.
Several recent reviews provide detailed coverage of several aspects of this topic. Particularly recommended are the general review of Maurizi (138), Gottesman and Maurizi on proteolytic regulation (79), Goldberg on ATP-dependent proteolysis (71), and Gottesman on minimizing proteolytic degradation of cloned proteins (76).
Lon (protease La). The Lon protease was the first ATP-dependent protease to be identified and characterized in detail. The enzyme was recognized biochemically by its ability to hydrolyze casein and a variety of other protein substrates (72). The protein was shown to be the product of the lon gene (29, 35), a locus at which mutations (originally called deg) that decrease the rate of degradation of certain fragments of β-galactosidase had been shown to occur (20, 81). Lon is a 350-kDa tetramer of 87-kDa subunits (71). The activity is inhibited by diisopropylfluorophosphate (DFP) (72), and a serine residue (Ser-679) believed to be at the active site has been identified (4). The enzyme is not a typical serine protease, however, and the deduced amino acid sequence (33) places Lon in a distinct class (188), along with other similar enzymes from eubacteria (109, 236, 237), yeasts (224), and humans (5, 249).
The Lon enzyme is extremely complex, with at least two enzymatic activities and at least four distinct binding sites on each subunit. Although important questions about the mechanism of action of Lon remain unanswered, a picture has emerged, mainly from the work of Goldberg, that succeeds in showing how the physiologically significant properties of the enzyme can be related to its mechanism of action (71, 79, 138). This mechanism requires distinct sites for peptide bond hydrolysis, for ATP binding and hydrolysis, and for the binding of protein substrates. The activity of the peptide bond-hydrolyzing site of Lon is regulated by two allosteric sites, one of which binds ATP and ATP analogs and the other of which binds proteins which are Lon substrates. Hydrolysis of small peptides is stimulated by both ATP and nonhydrolyzable ATP analogs. This indicates that in peptide bond hydrolysis, ATP functions not as a source of energy but, rather, as an allosteric effector, maintaining the peptide bond-hydrolyzing site in an open, active conformation. ADP strongly inhibits peptide bond hydrolysis and must therefore maintain the active site in a closed, inactive conformation. The enzyme has a higher affinity for ADP than for ATP (148, 149), suggesting that at the completion of each cycle of ATP hydrolysis the enzyme is left in an inactive state. Binding of a substrate protein (defined below) stimulates peptide bond hydrolysis directly (i.e., in the absence of ATP) and indirectly by stimulating the release of bound ADP and the binding of ATP. Thus, a substrate protein acts as an allosteric effector altering both the ATP binding site and the peptide hydrolysis site. Nonhydrolyzable ATP analogs do not support maximal rates of degradation for protein substrates (73). ATP must therefore play an additional role other than maintaining the active site in an open conformation. Although it is not clear how ATP hydrolysis allows Lon to degrade its macromolecular substrates, it is thought that energy is required to allow the substrate to be repositioned on the enzyme, exposing new cleavage sites (79). It is clear that Lon is a highly processive enzyme—the products of the degradation of protein substrates are 10- to 20-amino-acid peptides, and degradation intermediates are not produced even in the presence of a large excess of substrate (149). Thus, the substrate protein appears to remain attached to Lon during the degradation process. The hydrolysis of ATP appears not to be directly coupled to the hydrolysis of peptide bonds, since a Lon mutant lacking the active-site Ser and completely inactive as a protease retains wild-type levels of ATP hydrolysis (58). The ATPase activity of this mutant protein can be stimulated by substrate proteins as efficiently as the wild type.
Consideration of the specificity of Lon requires that we put aside the notion that protease specificity is determined solely by the amino acids forming or adjacent to the scissile bond. Although the peptide bonds cleaved frequently involve amino acids with aliphatic side chains (Leu or Ala), not all such bonds are actually cleaved, and a variety of other bonds may be cleaved (137). The major determinant of susceptibility to Lon cleavage is not the presence of a particular amino acid sequence at the cleavage site but, rather, the presence in the substrate protein of some signal that Lon recognizes. Although the nature of this signal is obscure, it is clear that it is absent from most native proteins but can be displayed when these proteins are denatured. The natural substrates of Lon (λN protein, RcsA, SulA, etc.) must display this signal normally. Since native proteins neither inhibit the hydrolysis of small peptides nor stimulate the release of ADP, the primary site at which Lon recognizes its substrates is not the peptide bond cleavage site but, rather, the protein binding site. It should be pointed out that other proteins may be involved in the recognition by Lon of certain abnormal proteins. As discussed below, heat shock proteins are required for the Lon-dependent degradation of certain abnormal proteins.
A particularly intriguing property of Lon is its ability to bind to DNA (36, 264). Binding to DNA stimulates both ATPase and protease activities. RNA will neither bind nor activate. It has been suggested that in the cell some of the Lon protease might be associated with DNA in order to degrade DNA-bound abnormal or regulatory proteins (71).
The biochemical properties of Lon seem to make sense for an enzyme designed to carry out the highly specific degradation of a special class of proteins in the presence of a very high concentration of other proteins without creating any deleterious products in the process. The activity of the enzyme is tightly regulated in that only when a protein displaying the recognition signal is bound to the substrate protein binding site is the peptide bond-hydrolyzing site activated. The absence of bound substrate protein would lead to the formation of the inactive ADP-bound form of the enzyme. Presumably, these elaborate mechanisms protect the normal proteins in the cell from Lon attack. The enzyme will not produce potentially deleterious protein fragments. Once a protein molecule is selected as a substrate, its degradation to small peptides is completed without the release of any intermediates.
The properties of lon mutants suggest that Lon plays two roles in the cell: it degrades a special class of proteins that are designed to be unstable (79), and it contributes to the degradation of other proteins that are damaged or in some way abnormal. Although lon null mutations are not lethal (141), strains carrying lon mutations show two distinct phenotypes: they are UV sensitive, and they form mucoid colonies. Both of these phenotypes are the direct result of the elevated levels of regulatory proteins (SulA and RcsA) that are normally degraded by Lon (79). In addition to degrading these naturally unstable proteins, Lon contributes to the degradation of abnormal proteins of various types. In lon mutants, the rate and extent of degradation of cell protein containing amino acid analogs or prematurely terminated by puromycin are reduced. This suggests that subsets of abnormal proteins display (or display when complexed to a heat shock protein) whatever signal Lon recognizes.
Consistent with its role in the degradation of damaged proteins, lon is part of the heat shock regulon (68, 183). Lon levels increase two- to threefold under conditions which induce this regulon. Overexpression of Lon from multicopy plasmids is deleterious (70), and as little as a fivefold increase in Lon levels results in a rapid cessation of growth.
Clp (protease Ti). ATP-stimulated degradation of casein can still be detected in extracts of E. coli strains carrying lon null mutations (114). The Clp protease (113) (protease Ti [104]) is responsible for much of this activity. This enzyme contains two dissimilar subunits: ClpP, responsible for peptide bond hydrolysis, and ClpA, an ATPase that is stimulated by protein substrates. ClpP is synthesized as a 21.7-kDa precursor which is autoprocessed by the removal of 14 N-terminal amino acids (139). Under high-salt (>0.1 M) conditions, ClpP exists as a 240-kDa dodecamer. The ClpA protein (84-kDa monomer) is purified as a dimer, which on exposure to Mg2+ and ATP is converted to a 500-kDa hexamer (138). Purified ClpA and ClpP associate in the presence of ATP to form a 750-kDa species thought to contain one ClpA hexamer and one ClpP dodecamer.
ClpP is a serine protease—it is inhibited by DFP (104), and both a serine and a histidine residue required for activity have been identified (140). The amino acid sequence indicates that ClpP is a member of a distinct structural class of serine proteases. Immunological evidence suggests that ClpP-like proteins are found throughout the biological world (140), and a number of genes encoding ClpP relatives found in plant chloroplasts have been sequenced (79). The amino acid sequence of ClpA indicates the presence of two distinct ATP binding sites (78). ClpA is a member of a large family of structurally related proteins (the ClpABC family) also found throughout the biological world (80, 213). It is not clear if all of these proteins function as components of proteolytic enzymes, and it has been suggested that the primary role of this family of proteins is to act as chaperones (180). In addition to ClpA, E. coli contains another member of the ClpABC family, ClpB (80). Another E. coli protein, ClpX (46 kDa) (77, 253), is a member of a related family (the ClpXY family). Both ClpA and ClpX but apparently not ClpB can associate with ClpP to form an active protease (ClpAP and ClpXP) (77). It is not clear how the Clp subunits are distributed among the possible enzymatic forms in the cell or whether this distribution is influenced by the presence of particular types of substrates.
Clp resembles Lon in that ATP is not required for the hydrolysis of small peptide substrates (254). The requirement for ATP in the hydrolysis of protein substrates is even more stringent for Clp than for Lon, however, and nonhydrolyzable analogs are inactive (104) in supporting hydrolysis of proteins. Clp shares several other important characteristics with Lon. Recognition of protein substrates depends not on interactions between the substrate and the peptide bond-hydrolyzing site but, rather, on interactions at a distinct substrate recognition site. In the case of Clp, the two sites are in different polypeptides. It is clear that the ATPase subunit determines the specificity of the Clp protease. ClpXP is able to hydrolyze the unstable λO protein but unable to hydrolyze casein. ClpAP is active on casein but is unable to hydrolyze λO protein (77, 253). As with Lon, it is not clear which features of the protein are recognized. Clp is processive: protein substrates are degraded to small peptides without the release of intermediates (138).
In vivo, ClpAP and ClpXP seem to be involved in the degradation of distinct classes of proteins. Mutants lacking clpA (113), clpP (139), or clpX (77) are not impaired for growth. The absence of either ClpA or ClpP leads to relatively minor effects on the degradation of canavanine-containing proteins and has no effect on protein degradation during starvation (113, 139). Mutations inactivating either ClpA or ClpP stabilize a ClpA-LacZ fusion protein as well as ClpA itself (78). Mutations in clpP or clpX stabilize not only λO protein but also a number of phage and cellular proteins (47, 66, 77, 202).
The clpP and clpX genes are cotranscribed as the clpPX operon (77). Transcription is elevated approximately twofold at high temperature, consistent with the observation that ClpP is a σ 32-dependent heat shock protein (123). clpA transcription is not dependent on σ 32 (113).
Alp. Although lon mutations increase the half-life of SulA and RcsA, these proteins are degraded more rapidly than most E. coli proteins even in lon mutants. A screen for E. coli genes that when overexpressed suppress the phenotypes associated with overproduction of SulA and RcsA in lon mutants led to the isolation of a plasmid carrying the alpA locus (alternate Lon protease) (238, 239). lon mutant strains carrying this plasmid show elevated rates of energy-dependent SulA degradation and presumably contain elevated levels of a new energy-dependent protease. It is clear, however, that this protease is not the product of the alpA locus. Instead, the presence of alpA on a high-copy-number plasmid activates transcription of the nearby slp (suppressor of Lon-like protease) gene, which in turn leads to the excision of a P4-like prophage which contains both alpA and slp (119). This excision inactivates a regulatory RNA molecule (10Sa RNA), encoded by the ssr gene, which overlaps one end of the cryptic prophage. It is this RNA which apparently acts as a negative regulator of the synthesis or activity of an energy-dependent protease which remains to be discovered.
Cytoplasmic Activities.
Protease II. This enzyme was discovered because of its ability to cleave chromogenic ester substrates of trypsin-like enzymes (171, 174). A gene encoding an enzyme with properties very similar to those of protease II has been cloned and sequenced (112). Although the molecular mass of the encoded protein (82 kDa) is different from that originally reported for protease II (58 kDa) (171), both reports describe a DFP- and N α-p-tosyl-l-lysine chloromethyl ketone (TLCK)-sensitive serine protease with strong specificity for classic trypsin substrates. The enzyme hydrolyzes peptide substrates (including the oxidized insulin B chain) at basic amino acid residues. Salmonella typhimurium (official designation, Salmonella enterica serovar Typhimurium) mutants (tlp) lacking protease II have been isolated and show no defect in any known proteolytic process (92). The deduced amino acid sequence indicates that in spite of its substrate specificity, this enzyme is not related to trypsin but is a member of the prolyl endopeptidase structural family (186).
Protease In. A cytoplasmic protease able to hydrolyze certain chromogenic peptide substrates for trypsin-like enzymes has been purified from E. coli (115). This enzyme is reported to be a 66-kDa monomer and to differ from protease II in both its sensitivity to inhibitors and its substrate specificity.
Protease Fa. Protease Fa has been identified biochemically as a cytoplasmic endoprotease able to degrade casein, globin, and certain N-terminal β-galactosidase fragments (72). The native enzyme has a molecular mass of ≈110 kDa and is inhibited by DFP and by metal chelators.
Protease So. Protease So has been identified biochemically as a cytoplasmic endoprotease able to hydrolyze casein, globin, glucagon, and denatured bovine serum albumin to acid-soluble peptides but unable to attack insulin or "auto-α," a small N-terminal fragment of β-galactosidase (37, 72). The enzyme is a dimer of 77-kDa subunits. Its sensitivity to DFP and resistance to metal chelators indicate that it is a serine protease. In vitro, protease Do selectively degrades oxidatively damaged glutamine synthetase (131) and cleaves the Ada protein (129).
Periplasmic and Membrane-Associated Activities.
"Protease I." Although the periplasmic protein "protease I" was first recognized by its ability to cleave chromogenic ester substrates of chymotrypsin-like proteases (175, 176), there has been considerable doubt that it has significant proteolytic activity (122). The nucleotide sequence of E. coli protease I has been determined (106) and is identical to that of a periplasmic thioesterase (34). Since it is clear that this enzyme can function in vivo as an efficient thioesterase and since no proteolytic defect has been associated with its loss (122, 153), it seems reasonable to discard the "protease I" designation for this enzyme.
Protease III (protease Pi, pitrilysin). The periplasmic metalloprotease protease III was discovered as an activity able to carry out the in vitro degradation of the α-complementing activity of certain N-terminal fragments of β-galactosidase (32). Because protease III is inactive on fragments larger than about 7 kDa and is able to degrade the oxidized B chain of insulin (30 amino acids), it was suggested that the enzyme is specific for small protein substrates (32). More recent studies suggest that protease III can hydrolyze a number of peptides in the 10- to 30-amino-acid size range but cannot attack smaller peptides or larger proteins (12 kDa and larger) (6). Other observations suggest that the action of protease III may not be restricted to small proteins, however, and that it may contribute to the degradation of secreted, protease-sensitive, large proteins (11). Protease III is periplasmic (although overproduction can result in significant release into the medium [51]) and is identical to the insulin-degrading enzyme protease Pi (72, 232, 233). Protease III is a metalloprotease, inhibited by chelators of divalent cations and reactivable by Zn2+, Co2+, or Mn2+ (32). The purified enzyme contains Zn2+ (52).
The gene encoding protease III, ptr, is located between recC and recB recD, with its 3' terminus overlapping the coding region for recB (57). The ptr gene can be transcribed from a promoter located between recC and ptr (39), and it is possible that transcripts from this promoter include recB and recD. There is also evidence for the existence of a larger transcript containing both ptr and all three rec genes (S. Kushner, personal communication). The deduced amino acid sequence predicts a protein of 108 kDa (57). This sequence places protease III in the insulinase structural family (3, 186), which includes human (3) and insect (126) insulinases and the small subunit of a mitochondrial processing protease (177). Nothing is known about the regulation of ptr or the physiological role of protease III.
DegP (HtrA, protease Do). DegP has been discovered by four independent routes: as a soluble protease that hydrolyzes casein (protease Do) (72, 200, 231), as an activity that participates in the degradation of abnormal periplasmic proteins (DegP) (216, 218), as a gene product required for growth of E. coli at high temperature (HtrA) (132, 134), and as a gene product required for S. typhimurium virulence (110). Although the enzyme was originally reported to be cytoplasmic (on the basis of its failure to be released on osmotic shock) (233), it has a signal peptide which is removed to form the mature protein (133, 134). DegP-alkaline phosphatase protein fusions express active alkaline phosphatase, suggesting that the N-terminal region of the protein can direct localization to the membrane or the periplasm (218). It is likely, therefore, that DegP is localized in the periplasm, although overexpression appears to result in accumulation of a cytoplasmic form of the enzyme (134). It is possible that the large size of the native enzyme interferes with its release from the periplasm by osmotic shock. The monomer molecular mass of the mature protein is 49 kDa (133). The native enzyme has an unusually high molecular mass: both a 500-kDa and a 300-kDa form are found, depending on growth conditions and strain background (231).
The purified protease degrades casein but does not attack a number of other proteins, nor does it hydrolyze any of the standard chromogenic ester substrates (134, 231). Sensitivity to DFP and resistance to other protease inhibitors place this enzyme in the serine protease family (134, 231).
Both the E. coli (133) and the S. typhimurium (110) degP (htrA) genes have been sequenced. The deduced amino acid sequences are very similar to each other and are closely related to the sequences of similar enzymes from Rochalimaea henselae (GenBank accession number L20127), Rickettsia tsutsugumushi (L11697), and Brucella abortus (L09274). An E. coli partial sequence ("htrH"; P31137) is very similar to degP (htrA), suggesting that a second member of this family is encoded in the E. coli genome. This group of sequences defines a new family of serine proteases (186).
It has been suggested that the major physiological role of DegP is to degrade abnormal proteins exposed to the periplasm (134, 216). Unstable membrane or periplasmic proteins are stabilized by degP mutations, but unstable cytoplasmic proteins are not (216). Several unstable periplasmic proteins have been found to be stabilized by degP (htrA) mutations (26, 159, 263). E. coli mutants lacking DegP are unable to grow at temperatures above 42°C (134, 200, 218), but the corresponding S. typhimurium mutants are not temperature sensitive (110). The E. coli growth defect is suppressed by lpp mutations, which allow periplasmic proteins to be released into the medium (216), and it has been suggested that the S. typhimurium strain in which mutations have been isolated may allow a higher rate of periplasmic leakage than E. coli (110). degP (htrA) is expressed at elevated levels at high temperature, and this elevated expression requires σ E (σ 24) rather than σ 32, the σ factor responsible for the transcription of most heat shock genes (55, 133).
SohB. Multicopy plasmids carrying the sohB (suppressor of htrA) locus suppress the temperature-sensitive phenotype of degP (htrA) mutations (9). The deduced amino acid sequence of SohB shows substantial similarity to that of protease IV (SppA), and it has been suggested that SohB may be a protease that, when overexpressed, can partially compensate for the loss of DegP (HtrA). Mature SohB (37 kDa) is derived from a 39-kDa precursor by removal of a signal peptide and is thought to be located in the periplasm. Null mutations of sohB cause no discernible phenotype.
Prc (Tsp, protease Re). Protease Prc has been identified by at least six independent routes: biochemically as a soluble protease able to degrade casein and other protein substrates (72, 179) and as an enzyme able to degrade oxidized glutamine synthetase (191), genetically and biochemically as a gene and enzyme responsible for the C-terminal processing of penicillin-binding protein 3 (88, 89) and of certain mutant λ repressor proteins (182, 206), and genetically as a gene conferring hypersensitivity to antibiotics (199) and as a gene affecting survival in macrophages (14). The nucleotide sequence (89, 205) predicts a precursor protein of 74 kDa. The native enzyme is a monomer. Although there is disagreement in the literature about the localization of the protein, it clearly contains a signal peptide which is processed to form the mature protein (205), and it seems likely that at least some of the enzyme is associated with the cytoplasmic membrane with the active site in the periplasm (89). When overexpressed, the enzyme is found in both the periplasmic and cytoplasmic membrane fractions (89, 205).
In several cases, Prc (Tsp) carries out the specific removal of a nonpolar C-terminal peptide (hence the names prc [processing C-terminal] and tsp [tail-specific protease]) (14, 89, 163, 205). The enzyme is apparently also capable of degrading casein and several other proteins to acid-soluble fragments (179). The activity is inhibited by DFP (although inactivation is unusually slow) and by N-tosyl-l-phenylalanine chloromethyl ketone (TPCK), suggesting that Prc (Tsp) is a serine enzyme (179). The amino acid sequence is not similar to that of any other known protease, although there is a striking similarity to regions of the bovine and human interphotoreceptor retinoid-binding protein (205).
In addition to the phenotypes noted above, prc (tsp) mutants are temperature sensitive for growth in low- but not high-salt medium (perhaps because of a defect in the heat induction of heat shock proteins) (89). prc (tsp) mutants also leak periplasmic proteins (89). It has been suggested that Prc (Tsp) functions in the degradation of proteins exposed to the periplasm, perhaps especially proteins damaged by osmotic or thermal stress (89).
Protease Mi. The periplasmic protease Mi activity has been purified from E. coli and shown to attack both casein and globin but not small N-terminal fragments of β-galactosidase (72). Its sensitivity to DFP indicates that it is a serine protease, but it is also sensitive to metal chelators and to dithiothreitol. The native enzyme has a molecular mass of about 110 kDa.
Protease IV (SppA). The cytoplasmic membrane protease IV activity was first recognized by its ability to hydrolyze chromogenic endoprotease substrates (172, 173). Later work identified protease IV as the major membrane-associated signal peptide peptidase activity on the basis of its ability to carry out the in vitro degradation of the major lipoprotein signal peptide (103, 105). Protease IV appears to be a tetramer of 67-kDa monomers (107) and is a serine protease (173).
Protease IV hydrolyzes synthetic chromogenic endoprotease substrates and has a preference for those containing hydrophobic amino acids (e.g., Z-Val, Phe, Leu, or Ala ONPs, where Z is N-benzyloxycarbonyl and ONP is p-nitrophenyl ester) (173). When a membrane preparation containing the lipoprotein signal peptide is exposed to protease IV, only the signal peptide appears to be degraded and there is no degradation of the other proteins present (105). Later workers found that although protease IV does not hydrolyze casein, it does produce acid-soluble fragments from a detergent-extracted mixture of E. coli membrane proteins (178), indicating that protease IV is able to hydrolyze substrates other than signal peptides. Protease IV digestion of the isolated lipoprotein signal peptide (in the absence of cell membranes) results in the production of a number of small peptide products (169). The identity of these products suggests that protease IV does not require a free N or C terminus, prefers hydrophobic amino acids on either side of the scissile bond, and will not cleave a peptide containing fewer than six amino acids. All but one of the protease IV cuts occurred in the hydrophobic region of the lipoprotein signal peptide. It is not clear why protease IV is unable to recognize the signal peptide before it is cleaved from the precursor. Perhaps the signal peptide has a different conformation after its release from the precursor (169). Neither overexpression nor loss of protease IV by mutation is deleterious to cell growth. Null mutations in sppA do not lead to the detectable accumulation of signal peptides (230).
The gene encoding protease IV (sppA [signal peptide peptidase]) has been cloned and sequenced (107). The deduced amino acid sequence shows significant similarities to two recently described enzymes: a "signal peptidase" from Leptospira borgpetersenii (GenBank accession number L27482) and the product of the E. coli sohB gene (see above) (9).
Proteases V and VI. The cytoplasmic membrane proteases V and VI are less well characterized than protease IV. Protease V can be purified from detergent extracts of E. coli membranes as an activity that hydrolyzes chromogenic ester substrates (173). Protease V preferentially hydrolyzes N-blocked p-nitrophenyl esters of Gly, Leu, and Val, distinguishing it from protease IV (SppA), which prefers the corresponding Ala and Leu compounds. The purified enzyme is reported to hydrolyze radiolabelled E. coli cytoplasmic proteins (173) and to hydrolyze casein (178). Both proteases V and VI are serine proteases. It appears that the enzyme described by Regnier as protease IV corresponds to protease V (178, 187).
A third cytoplasmic membrane protease, protease VI, has been isolated on the basis of its ability to hydrolyze a mixture of radiolabelled E. coli membrane proteins (178). Although protease VI is also a DFP-sensitive serine protease, it differs from proteases IV and V by its inability to hydrolyze chromogenic ester substrates and its sensitivity to benzamidine, an inhibitor of trypsin-like enzymes. No mutants lacking either protease V or protease VI have been described, and the genes encoding them have not been cloned.
HflA and HflB. The level of the proteolytically unstable bacteriophage λ cII protein is thought to determine whether infection will proceed via the lytic or the lysogenic pathway (99). Mutations at the hflA (high frequency of lysogenization) locus increase the frequency of lysogeny by stabilizing cII. The hflA locus contains three genes: hflX, hflK, and hflC (12, 168), and the products of two of these (HflK and HflC) are present in the purified protease (31). The HflA protease appears to be associated with the cytoplasmic membrane (31, 168, 266), although it is likely that its active site is in the cytoplasm (168). Purified HflA has a molecular mass of approximately 200 kDa and is thought to be composed of two HflC, one HflK, and one HflK', a proteolyzed form of HflK. The enzyme is sensitive to phenylmethylsulfonyl fluoride and is likely to be a serine protease, although it is also sensitive to divalent cations, especially Zn2+ and Cu2+ (31). The purified enzyme hydrolyzes the cII protein (11 kDa) to acid-soluble fragments (31), but the rate of degradation is relatively low. On the basis of sequence similarity, HflX appears to be a GTP-binding protein, and it is possible that HflX is required for full proteolytic activity (168). HflC contains a region with amino acid sequence similarity to a segment of ClpP that includes the catalytic serine residue (168).
The function of the HflA protease in the cell is unknown. Although mutants containing hflA null mutations are viable (13), the levels of a number of cellular proteins are elevated in hflA mutants (30). Some of these proteins are unstable and may be cellular substrates for the HflA protease. It is thought that the effects of the cyclic AMP (cAMP)-cAMP receptor protein system on the lysis/lysogeny decision are mediated by HflA, and it is interesting that several of the proteins that show elevated levels in an hflA mutant also have elevated levels in cya mutants grown in medium containing cAMP.
The cII protein is degraded much more rapidly than most proteins even in hflA mutants (102), indicating that there is an HflA-independent pathway for cII degradation. Mutations at the hflB locus also increase the frequency of lysogenization by λ (13), and it has been suggested that hflB might encode a component of a protease distinct from HflA (13). The hflB gene is identical to ftsH, a gene required for growth at high temperature (98). The ftsH (hflB) gene encodes a cytoplasmic membrane protein with strong sequence similarity to a family of eukaryotic ATPases. It has been suggested that this protein is either a component of a proteolytic system or a chaperone which presents cII to a protease.
OmpT (protease VII) and related enzymes. OmpT is an outer membrane serine protease highly specific for dibasic sites (e.g., -Arg-Arg-) (222). The purified enzyme is sensitive to DFP and to certain divalent cations (Cu2+, Zn2+, and Fe2+) (223). The specificity of OmpT for paired basic residues is quite rigid (although there are reports of cleavage at -Arg-Met- [265] and -Arg-Ala- [128] bonds), and the enzyme will not attack the standard chromogenic trypsin substrates which contain a single basic amino acid. The enzyme is an endoprotease and will not cleave the bond between two basic amino acids if one of them is at either the C or N terminus of the peptide. These specificity properties are quite different from the paradigmatic basic-residue-cleaving protease trypsin, and OmpT is not a trypsinlike enzyme.
The deduced amino acid sequence of OmpT predicts a mature protein of 33 kDa (83, 223). The purified, detergent-solubilized enzyme has a molecular mass of about 180 kDa (223). The enzyme is very similar to another E. coli protease, OmpP (116), and to related enzymes from S. typhimurium (protein e [82, 262]) and Yersinia pestis (plasminogen activator/coagulase [145, 211]). This group of enzymes forms a distinct class of serine proteases (186).
Although the physiological function of OmpT is not clear, it is clear that OmpT can contribute to the degradation of proteins that can come into contact with the outer membrane. The TonB protein, required for active transport of vitamin B12 and iron siderophores across the outer membrane (111, 185), is proteolytically unstable in the absence of an associated protein, ExbB (209). This instability depends on OmpT: ompT exbB mutants contain normal levels of the TonB protein (208). This observation suggests that OmpT contributes to the degradation of periplasmic proteins, which is consistent with the observed OmpT-dependent degradation of certain periplasmic fusion proteins (10). OmpT levels are regulated by growth temperature, with very low expression at 30°C and much higher levels at 42°C (192).
The proteolytic inactivation of the Shigella flexneri VirG protein expressed in E. coli provides another example of an in vivo process involving OmpT (166). VirG is an outer membrane protein required for the inter- and intracellular spread of Shigella and enteroinvasive E. coli strains in epithelial cells. When VirG is expressed in E. coli K-12, only a fragment of the VirG protein is found in the outer membrane and the cells are unable to spread. Mutants of K-12 lacking OmpT express active VirG and are able to spread. Shigella and enteroinvasive E. coli strains lack OmpT. These results indicate that OmpT can play an important role in determining the nature of the proteins within or in contact with the outer membrane.
It is quite clear that OmpT is responsible for a large number of proteolytic artifacts. Many of these have been shown to result from exposure of susceptible proteins to OmpT during cell lysis or subsequent purification, but others arise as a result of in vivo OmpT action on proteins that can contact the outer membrane. Examples of normal bacterial or phage proteins attacked by OmpT include the ferric enterobactin receptor (59, 60, 101), phage T7 RNA polymerase (83) (which is also attacked by S. typhimurium protein e [82]), the UvrB protein (24), the ω subunit of E. coli RNA polymerase (64), E. coli initiation factor IF2 (128), E. coli penicillin-binding proteins 7 and 1b (93), and the Ada protein (198). OmpT can associate with particles of phage φX174 and cleave the prohead accessory protein gpB (189), but this association and cleavage are probably not physiological (46). A number of foreign proteins expressed in E. coli undergo OmpT-dependent cleavage (17, 27, 91). In many cases, fusion proteins designed to allow secretion to the periplasm or to the outside of the cell seem to be particularly sensitive to OmpT cleavage (10, 86, 87, 246).
OrfX (glycoprotease?). An open reading frame (orfX) divergently transcribed from the rpsU dnaG rpoD operon of both E. coli (167) and S. typhimurium (54) codes for a protein with an amino acid sequence 76% identical to that of a glycoprotease from Pasteurella haemolytica A1 (1). The Pasteurella enzyme is a metalloprotease that is highly specific for O-glycosylated glycoproteins and will not cleave proteins without this modification. The Pasteurella enzyme can be expressed in E. coli, where it is exported to the periplasm without cleavage of any amino acids from its N terminus. The promoter for the E. coli orfX gene has been shown to be functional (167).
Signal (leader) peptidases. Two proteases responsible for the cleavage of signal peptides from precursor proteins are localized to the cytoplasmic membrane. One of these enzymes, signal peptidase II (encoded by the lsp gene), is an 18-kDa enzyme that specifically cleaves signal peptides from glyceride-modified prolipoproteins (see, e.g., reference 162). The other enzyme, signal peptidase I (encoded by the lep gene), is a 37-kDa protein that acts on all other signal peptide-containing preproteins (reviewed in references 44 and 45). Both proteins are integral membrane proteins with periplasmic active sites. These two enzymes appear to be structurally unrelated. Signal protease I, although not inhibited by any of the classical protease inhibitors, seems to have a serine at the active site, and the group of related proteases of which it is the prototype constitutes a new family of serine hydrolases (45). The possibility that E. coli contains a third signal peptidase is suggested by the existence of an open reading frame (hopD) (100, 250) with sequence similarity to pilD, a gene encoding a peptidase/methylase that acts on the prepilin protein of type 4 fimbriae of Pseudomonas aeruginosa (221).
The enzymes discussed in this section are all able to attack small (2- to 5-amino-acid) peptides. It does not necessarily follow that they are capable of hydrolyzing only small peptides or that this is their only role in the cell. Unless otherwise noted, all of the enzymes discussed are present in both E. coli and S. typhimurium.
Dipeptidases.
Although dipeptides may be hydrolyzed by enzymes that can also attack larger peptides (all of the "aminopeptidases" show at least some activity toward dipeptides), several enzymes can hydrolyze only dipeptides.
Peptidase D. Peptidase D is a broad-specificity dipeptidase able to hydrolyze almost any dipeptide except those containing C-terminal Pro. Although PepD is the only enzyme in the cell that hydrolyzes carnosine (β-alanyl-l-histidine) (120), it also hydrolyzes many other peptides, and the designation of the activity as an "X-His dipeptidase" or "aminoacyl histidine dipeptidase" found in database entries is misleading. The nucleotide sequence of the E. coli enzyme has been determined (94, 96, 97, 121). The deduced amino acid sequence is not similar to that of any known protein. pepD transcription increases (~fivefold) in response to phosphate limitation, but this regulation is independent of the regulatory genes of the pho regulon (94).
Peptidase Q. Peptidase Q specifically hydrolyzes X-Pro dipeptides (147). The E. coli enzyme has been sequenced (164), and the deduced amino acid sequence is quite similar to that of a human enzyme (prolidase or peptidase D) with the same specificity (234). It is interesting that the mutational loss of this enzyme confers a similar phenotype at the molecular level in both bacteria and humans: X-Pro dipeptides accumulate (53, 152). A strain lacking both peptidases P and Q is unable to utilize X-Pro peptides and cannot fully degrade intracellular proteins (152), accumulating small Pro peptides.
Peptidase E. Peptidase E is the only one of the three S. typhimurium Asp-X-specific peptidases (25) to be characterized in detail. No other enzymes with this specificity have been described. The S. typhimurium gene has been cloned and sequenced, and its product, peptidase E, has been crystallized (41). The sequence of E. coli pepE has also been determined (18). The enzyme is not inhibited by any of the standard types of inhibitors, and its deduced amino acid sequence is unrelated to that of any known protein. Transcription of the pepE gene is regulated by Crp.
Other dipeptidases. A specific β-alanyl dipeptidase has been purified from E. coli (84). This enzyme must be different from PepD, which can also hydrolyze β-alanyl peptides, since PepD also hydrolyzes many other peptides whereas the β-alanyl dipeptidase is active only toward β-alanyl peptides. An enzyme able to hydrolyze Gly-Gly has been identified in extracts of a pepN pepA pepB pepD pepP pepQ S. typhimurium strain, and mutations (pepG) lacking the activity have been isolated (C. Miller, unpublished data). There are therefore at least seven distinct dipeptidases in E. coli and S. typhimurium (D, E, Q, G, two additional Asp-X hydrolyzers, and the β-alanyl peptidase).
Tripeptidase.
Peptidase T. Peptidase T is an aminotripeptidase. Although the substrate specificity of PepT has not been thoroughly characterized, it appears to be quite specific for tripeptides but rather nonspecific for the particular amino acids in the peptide (217, 219). The S. typhimurium enzyme has been cloned and sequenced, and there is also a partial sequence for E. coli pepT (62, 135). Identification of a PepT homolog (47% amino acid sequence identity) in Lactococcus lactis (150) suggests that this type of enzyme may be present in many bacterial species. The deduced amino acid sequence of pepT shows some similarity to carboxypeptidase G2, a Pseudomonas enzyme that removes glutamate from folic acid and its derivatives and analogs (150, 156, 158).
The pepT gene is transcribed from two promoters: synthesis from p1 is increased in anaerobiosis, and this regulation depends on Fnr, a transcriptional regulator required for elevated anaerobic expression of many anaerobic genes, especially those whose products play a role in anaerobic respiratory pathways (212). Transcription from the upstream p2 promoter appears to be constitutive, providing a low level of pepT expression under aerobic conditions (156). The p2 promoter overlaps the promoter for the divergently transcribed potABCD operon, which encodes a putrescine/spermidine transport system (62, 135). The physiological significance of the anaerobic regulation of pepT is unclear. It seems possible that PepT makes available amino acids that could participate in anaerobic respiratory pathways.
Peptidase A. Peptidase A is a broad-specificity aminopeptidase that is able to remove many N-terminal amino acids from peptides in which the amino acid adjacent to the N terminus is not proline (155, 245). The deduced amino acid sequence of pepA shows that it is a member of the leucine aminopeptidase (LAP) family (215). A crystal structure is available for one member of this family, bovine lens leucine aminopeptidase (BLAP) (21, 22, 23, 118). BLAP contains six identical 54-kDa subunits and 12 Zn2+ ions. Comparison of the sequences of BLAP and PepA indicates that all the important structural features of the active site are conserved between the two enzymes and that all the amino acid residues involved in metal ion binding are absolutely conserved in Pep A (22).
E. coli pepA mutants are defective in the site-specific recombination system that resolves plasmid multimers in ColE1 plasmids (215). This system, necessary for the stable maintenance of ColE1, requires a site on the plasmid (cer) and two chromosomally encoded genes: xerA and xerB. The sequence of the xerB locus revealed similarity to BLAP, and further analysis led to the conclusion that xerB is pepA. Although the mechanism by which PepA participates in site-specific recombination is not understood, it is clear that the peptidase activity of PepA is not required for its role in site-specific recombination (144).
Peptidase B. Peptidase B is another broad-specificity aminopeptidase. The deduced amino acid sequence of PepB shows that it, too, is a member of the LAP family (Z. Mathew and G. G. Miller, unpublished results).
Peptidase N. Peptidase N, a third broad-specificity metalloaminopeptidase, is exclusively responsible for the hydrolysis of certain chromogenic peptidase substrates (e.g., Ala β-naphthylamide), which can be used to detect the enzyme in colonies growing on an agar surface. The deduced amino acid sequence (61, 143) shows that PepN is a member of the large family of alanyl aminopeptidases (186), which includes bacterial, fungal, and mammalian enzymes. Many of the mammalian enzymes are membrane-associated glycoproteins thought to play important roles in the hydrolysis of peptides in brush border membranes of the small intestine, renal proximal tubules, synaptic membranes of the nervous system, and macrophage surfaces. Although there were early indications that E. coli peptidase N was in some way associated with the membrane, nothing in the amino acid sequence suggests such an association, and the most closely related bacterial PepN (from Lactococcus lactis) has been shown to be cytoplasmic (243).
The pepN gene has been reported to be transcriptionally regulated by anaerobiosis and by phosphate limitation (67). The anaerobic induction of pepN could not be observed in fusions of the pepN promoter to reporter genes (61). Mutations in fnr, crp, phoB, phoM, or phoR did not seem to affect pepN expression (67).
Peptidase P. Peptidase P is an aminopeptidase that specifically removes N-terminal amino acids adjacent to a proline residue (256). The deduced amino acid sequence of PepP (260, 261) shows that it is related to PepQ and to PepM. Each of these peptidases shows strong specificity for the amino acid contributing the nitrogen atom of the cleaved bond, and proline is accepted (PepM) or required (PepP, PepQ) in this position. It is likely that the three-dimensional structures of these three peptidases are even more similar than the amino acid sequences would indicate (15).
Iap. It has been known for over 30 years that E. coli produces several electrophoretically separable forms of alkaline phosphatase (AP) (reviewed in reference 165). These forms result from the partial removal of a single Arg residue from the N terminus of the mature AP. Since AP is a dimer, this partial modification results in three forms of the enzyme: homodimeric unmodified, heterodimeric, and homodimeric modified. Mutants unable to carry out this modification carry lesions at the iap locus, which codes for a membrane-associated protein which has been cloned and sequenced (108). Although the sequence in its entirety does not resemble any other known protein, it contains a subsequence of about 45 amino acids with significant similarity to PepT, PepD, and Pseudomonas carboxypeptidase G2 (156).
It seems likely that iap codes for an N-terminal Arg-specific aminopeptidase. When a chimeric protein containing the E. coli AP signal peptide joined by an Arg residue to a mammalian cytochrome b 5 is expressed in E. coli, the Arg residue is removed after signal peptidase cleavage by a metal chelator-sensitive aminopeptidase (90). The same conditions that inhibit modification of the chimeric AP-Arg-cytochrome b 5 inhibit formation of AP isozymes, suggesting that both are removed by the same enzyme. Although it has not been rigorously shown that this enzyme is the product of the iap gene, it seems likely that it is. It is thought that the modification of AP is not physiologically significant, and the role of this enzyme is unknown (108).
γ-Glutamyl transpeptidase. γ-Glutamyl transpeptidase is named for its transpeptidase activity, but its periplasmic location and its greater γ-glutamyl peptidase than transpeptidase activity suggest strongly that its major cellular function is in catabolism (225). Mutants with mutations in the gene encoding this enzyme (ggt) have been isolated (229), and these mutants are unable to use γ-glutamyl peptides (e.g., glutathione) as sources of amino acids. γ-Glutamyl peptides cannot be transported into the cell and require hydrolysis in the periplasm before they can be catabolized (225). This contrasts with the utilization pathway for most small peptides, which are transported into the cytoplasm and hydrolyzed there. The gene has been cloned and sequenced (226, 227), and the protein has been purified and crystallized (125). Synthesis of Ggt is temperature dependent: the level is high at 20°C, intermediate at 37°C, and below detectability at 43°C (228).
Peptidase M and N-terminal Met removal. In bacteria, the synthesis of all known proteins is initiated with N-formyl Met. Nearly all proteins are processed by a specific deformylase (2, 142) to remove the N-formyl group, and many proteins are further processed to remove N-terminal Met. The importance of the amino acid adjacent to the N-terminal Met in determining whether processing occurs was predicted by early studies showing that the distribution of N termini in E. coli proteins is highly nonrandom (194, 248) and by more recent comparisons of N-terminal protein sequences with the predictions of the corresponding nucleotide sequence (242). These studies led to the conclusion that N-terminal Met can be removed only if the adjacent amino acid is small; Ala, Pro, Ser, Thr, Gly, Cys, and Val are permissive, whereas other amino acids are nonpermissive (203). Interestingly, the rules for N-terminal Met removal are the same for all organisms that have been studied (7, 203).
Although the existence of N-terminal Met removal has been recognized for many years, the genes and enzymes involved have only been found in the last few years. The aminopeptidase responsible for removal of methionine is the product of the pepM gene, which has been cloned and sequenced from both S. typhimurium (157, 161) and E. coli (16). The product of the gene, PepM or methionine aminopeptidase, possesses enzymatic properties which are entirely consistent with a role in N-terminal Met removal: it is highly specific for N-terminal Met, it is exquisitely sensitive to the nature of the amino acid adjacent to the N-terminal Met, and it normally removes only a single amino acid from the N terminus of its substrates (16, 252). PepM removes Met from N-terminal Met tripeptides if the residue adjacent to the N terminus is one of the permissive amino acids listed above. The specificity for the N-terminal amino acid is high but not absolute: both Met sulfone and Met sulfoxide can be cleaved at a detectable rate by the Salmonella enzyme, and peptides containing the Met analog norleucine at their N termini are also cleaved (J. L. Miller, unpublished observations). The enzyme is completely inactive on N-blocked substrates (e.g., N-formyl Met), has a very low activity with certain Met dipeptides, and is capable of removing N-terminal Met posttranslationally from some proteins (e.g., recombinant interleukin-1β [157]) but not others (J. L. Miller, unpublished observations).
PepM is a metallopeptidase that is maximally active in the presence of Co2+, although the identity of the metal ion bound in the cell has not been determined (252). The X-ray structure of PepM (190) reveals a novel active-site structure that is quite different from that of LAP or carboxypeptidase. The "pita bread fold" that forms the active site of PepM is also present in a creatinase from Pseudomonas putida and is likely to be present in several other enzymes (including PepP and PepQ) (15).
pepM is a vital gene in both S. typhimurium (154) and E. coli (28) and is the only one of the known peptidase genes essential for viability. Nothing is known about the regulation of pepM.
Carboxypeptidase.
Dipeptidylcarboxypeptidase. The only C-terminal exopeptidase known to be present in E. coli or S. typhimurium removes dipeptides from the C termini of its substrates (255, 257). The enzyme requires a free C terminus, and it cannot hydrolyze bonds in which the peptide nitrogen is donated by proline or in which both amino acids are Gly. The dcp gene has been cloned and sequenced from both S. typhimurium (85) and E. coli (95). The deduced amino acid sequence shows strong similarity to that of oligopeptidase A and the other members of the "thimet" family of metallopeptidases (186).
Although Dcp can function as a catabolic enzyme (it is required for the utilization of N-acetyl Ala3 (AcAla3) as the sole nitrogen source [50, 243a]), it seems unlikely that catabolism is its major function. N-blocked peptides are poorly transported into the cell, and most unblocked substrates of Dcp (four amino acids or larger) would also be expected to be transported poorly. It is more likely that the major cellular function for Dcp is in the degradation of intracellular proteins, discussed below.
Other Peptidases.
Oligopeptidase A. OpdA was originally detected as an activity able to hydrolyze AcAla4 in extracts of a dcp−S. typhimurium strain (244). OpdA hydrolyzes N-blocked peptides with at least four amino acids (e.g., AcAla4), while at least five amino acids are required for an unblocked peptide to be hydrolyzed (e.g., Ala5). The ability of a substrate to be hydrolyzed is influenced by amino acids other than those contributing to the scissile bond, since N-benzyloxycarbonyl (Z)-Gly4 and Z-Gly5 are not substrates whereas Z-Gly-Pro-Gly-Gly-Pro-Ala is hydrolyzed at the Gly-Gly bond. Ala or Gly on either side of the scissile bond is permissive for hydrolysis, but the number of peptides tested is not sufficient to determine the range of allowable amino acids.
In vitro OpdA is the major soluble activity able to hydrolyze the prolipoprotein signal peptide (169, 170). Analysis of the products of OpdA-catalyzed hydrolysis of the 20-amino-acid signal peptide revealed six cleavage sites, each involving either Gly or Ala on one side or the other of the scissile bond. OpdA did not attack the prolipoprotein itself, suggesting that it recognizes an altered conformation of the peptide achieved only after release from the precursor. An alternative hypothesis, i.e., that the enzyme is able to attack only peptides below a certain minimum size, appears to be excluded by recent studies of the OpdA-catalyzed hydrolysis of the bacteriophage P22 gp7 protein.
Mutants of S. typhimurium lacking OpdA are unable to support normal growth of bacteriophage P22 (43). In the wild-type cell, the phage protein gp7 is produced by the removal of 20 amino acids from the N terminus of its 229-amino-acid precursor. This processing does not take place in an opdA mutant. No other phage or bacterial protein is required for the processing reaction, which can be observed in vitro with purified OpdA (A. Toguchi and C. Conlin, unpublished results). The bond apparently cleaved by OpdA (Glu-Lys) does not fit the rules deduced from either the small-peptide studies or the prolipoprotein signal peptide cleavage studies. Although none of the substrates previously tested contained such a bond, it is tempting to speculate that OpdA recognizes something other than the nature of the amino acids forming the scissile bond.
Both the S. typhimurium (40) and E. coli (42) versions of the gene encoding OpdA have been cloned and sequenced. The amino acid sequence deduced for OpdA is quite similar to that of Dcp. Both proteins are 680 amino acids in length and share (comparing the two Salmonella proteins) 33% identical amino acids along their entire lengths. One region, apparently a Zn2+-binding site, is particularly strongly conserved. The primary sequence similarity between Dcp and OpdA is greater than that between any of the other E. coli or S. typhimurium peptidases for which sequence information is available. The amino acid sequences of Dcp and OpdA also show striking similarities to a group of proteases found in a variety of organisms, with examples from eubacteria, fungi, and animals. Although the sequence similarity is particularly striking in the vicinity of the putative Zn2+-binding site there is clear similarity over large regions of the sequence. Three of these enzymes (OpdA, rat metalloendopeptidase, and rat mitochondrial intermediate peptidase) are known or thought to play a role in protein processing.
The opdA sequence 5' to the coding region contains a near-consensus σ 32 recognition site (40). Primer extension analysis has shown that this promoter is actually used, and results with mutants defective in σ 32 (htpR) have shown that σ 32 is required for opdA transcription (C. A. Conlin, unpublished data). The opdA gene is therefore a member of the heat shock regulon. The dcp gene shows no heat shock promoter, and there is no evidence that it is regulated.
The results of in vitro studies suggest that OpdA may play a special role in the degradation of signal peptides. Signal peptides are normally degraded so rapidly that they can almost never be observed in vivo. The major cytoplasmic activity able to degrade one such signal peptide (lipoprotein) is OpdA (170). A further indication that OpdA interacts in some special way with signal peptides comes from the discovery that prlC mutations are alleles of opdA (42). prlC mutations suppress the localization defect conferred by certain mutations in the signal sequence of the LamB protein (240, 241). Although the basis of the suppressor phenotype is not understood, the isolation of such mutations in opdA suggests that the protein can be mutationally altered to allow it to recognize the signal peptide before it is released from the precursor.
Metalloendopeptidase QG. A cytoplasmic metallopeptidase with a preference for -Gln-Gly- (QG) bonds has been purified from extracts of E. coli (184). This activity cleaves various small peptides (up to a nonapeptide) containing an internal Gln-Gly bond and is inactive on the oxidized A and B chains of insulin, which do not contain such a bond. The enzyme is a 67-kDa monomer, similar to Dcp and OpdA, which are also metallopeptidases, but neither its specificity nor its chromatographic properties suggest that it is identical to one of these enzymes.
Ecotin, the product of the eco (eti) gene (56, 130), is a 38-kDa homodimeric protein present in the periplasm of E. coli (38). Ecotin is a potent inhibitor of a variety of mammalian serine proteases including trypsin, chymotrypsin, elastase, and coagulation factor Xa (38, 146, 201). Ecotin differs from the classical mammalian serine protease inhibitors in its broad specificity, and it appears to be structurally unrelated to these proteins (130, 146). The physiological function of Ecotin is unclear. None of the known E. coli proteases is sensitive to Ecotin inhibition (38). It is thought that Ecotin may protect against proteolytic damage that might be caused by exposure to proteases in the host.
Several phage proteins are known to inhibit the degradation of unstable phage proteins. The product of the T4 pinA gene appears to be a relatively specific inhibitor of Lon (210). Expression of the PinA protein in growing cells leads to a Lon-like phenotype but does not inhibit Lon-independent proteolytic processes. λ cIII diminishes the rate of cII degradation, perhaps by inhibiting the HflA protease (31). cIII also inhibits the degradation of σ 32 (8), which is degraded by an unknown protease different from HflA. It has been proposed that the λ RexB protein may act to decrease the rate of λO degradation by inhibiting the protease (now known to be ClpXP) responsible for λO degradation (196).
It is useful to distinguish two functions for protein degradation in growing cells (138). Some substrates are degraded to rid the cell of damaged or nonfunctional proteins or peptides (housekeeping degradation); for other proteins, degradation is used to regulate the level of a functional protein (regulatory degradation). In starving cells, functional (although perhaps not functioning) proteins that are not normally degraded become susceptible to degradation. Note that this classification does not imply that specific protein degradation pathways are exclusively dedicated to each type of degradative process.
A few normal cellular processes are known to generate substrates for housekeeping degradation. Signal peptides cleaved from secreted proteins are degraded so rapidly that it is difficult to detect them (151a). Peptides synthesized from the leader regions of mRNAs for attenuation-controlled operons are also so rapidly degraded as to be essentially undetectable in vivo (48). In a few cases, irreversible inactivation occurs when a protein carries out its normal function. The Ada protein, for example, is irreversibly alkylated as it participates in the repair of alkylated DNA (193). The inactivated protein is then degraded. Many other processes occurring in growing cells may generate substrates requiring degradation. Translational errors (including not only amino acid substitutions but also premature termination and internal out-of-frame initiation) seem likely to lead to the formation of degradation substrates. Errors in folding which generate irretrievably misfolded products may also generate polypeptides that are degraded. Transient imbalances in the concentrations of proteins that normally associate to form complexes would presumably also contribute to the pool of rapidly degraded proteins. At least 20% of the peptide bonds synthesized in normally growing cells are rapidly hydrolyzed (259), and it seems likely that all of these processes may contribute to the pool of substrates for this rapid degradation. Other processes may damage normally stable proteins, converting them into degradable, abnormal proteins. A surprisingly large number of potential pathways, many involving normal cell metabolites, can lead to protein damage that might generate hyperdegradable proteins (214). Unfortunately, we do not know to what extent any of these processes contribute to the flux of proteins through the degradation machinery, and we cannot specify how various stresses affect this flux.
Starvation for any of a variety of required nutrients leads to an elevated rate of degradation of proteins that are stable in growing cells (151). Little progress has been made toward understanding the basis of the increased susceptibility of these proteins to degradation, the extent to which the degradation is selective, or the enzymes that are responsible.
Pathway of Protein Degradation.
Although it is not possible to specify the pathway by which any particular protein is broken down into its constituent amino acids, the properties of both the Lon and Clp proteases suggest that the process does not involve large peptides as intermediates. Both of these enzymes generate relatively small (<10-amino-acid) peptides. It is a reasonable guess that the unknown enzyme(s) that carries out starvation degradation is similar to Lon and Clp in this respect. Enzymes that hydrolyze these peptides to free amino acids are therefore necessary to complete the degradation process. It is clear that some of the same enzymes that participate in peptide catabolism are also required for the complete breakdown of virtually all types of degradable intracellular proteins (152, 258, 259). Thus, enzymes that are required for the utilization of small peptides as sources of amino acids are also required for degradation of intracellular proteins. In mutants lacking peptidases, intracellular proteins are not broken down to free amino acids, and a heterogeneous mixture of small peptides accumulates instead.
Several important cellular regulatory processes involve rapidly degraded proteins as key elements. Since proteolytic regulation has been the subject of a recent review (79), only the basic principles and a few examples will be discussed here.
For a protein to function in regulation, either its activity or its level (or both) must respond to a regulatory signal. For the paradigmatic regulatory proteins (lac repressor or Crp, for example), the activity of the regulator is affected, usually as the result of a conformation change induced by binding an effector. In some cases, however, the level of the regulator responds to the signal, and in a few cases the level of the regulator is determined mainly by its susceptibility to proteolysis. The regulatory protein may be unstable under normal growth conditions ("constitutively unstable" [75]) and respond to a regulatory signal that inhibits its degradation or, most often, elevates its rate of synthesis above the capacity of the degradation apparatus. There are several potential advantages inherent in this regulatory strategy. A protein can be maintained at a very low level if it is degraded rapidly. When its rate of synthesis is increased, the amount of the regulatory protein can increase dramatically. When the protein is no longer needed and its rate of synthesis decreases, proteolysis leads to its rapid destruction, restoring normal levels without the need for dilution by growth. As a result, an effective concentration of the regulatory protein may be present for only a short time. Several examples of this regulatory strategy have been studied. The SulA protein is a cell division inhibitor produced as part of the SOS response to DNA damage. SulA is degraded by Lon with a half-life of approximately 1 min, and it is present in normally growing cells at an undetectably low level. In response to DNA damage, transcription of sulA is derepressed and SulA levels rise to an effective level (160). When repression of sulA is restored, SulA levels are rapidly returned to their normal, low levels as a result of proteolysis. A similar sort of mechanism may be responsible for spatially limiting the action of certain proteins. Transposases encoded by transposable elements are frequently cis acting: they are efficient catalysts only of reactions involving the element that encodes them and are unable to act on other identical elements in the cell. In some cases, this limitation has been shown to result from the extreme proteolytic instability of the transposase protein (49, 63). These examples illustrate the use of proteolytic regulatory strategies for limiting the action of a protein in time (SulA) and in space (transposase). In other cases, the regulatory protein is normally stable and becomes unstable only in response to a regulatory signal. The classic example of such a strategy also comes from the SOS regulon. The LexA protein, the repressor of the genes of the SOS regulon, is stable in normally growing cells. In response to DNA damage, a signal is produced which activates the RecA protein to bind to LexA. This binding, in turn, activates the autoproteolytic activity of LexA, leading to cleavage and inactivation of the LexA protein (247). A more complex example is provided by the σ 32 protein, the σ factor responsible for transcription of the genes of the heat shock regulon (65). σ 32 is extremely unstable to proteolysis under normal growth conditions. This instability depends on DnaK, and it is thought that the complex of DnaK with σ 32 is much more unstable than is free σ 32. Under conditions that induce the heat shock regulon, DnaK is attracted to other proteins, leaving σ 32 uncomplexed and therefore relatively stable.
It is important to note that we have no idea how many rapidly degraded proteins there are. If the cell makes use of this strategy as frequently as phage λ, which encodes at least six unstable regulatory proteins (see below), there must be many more to be discovered. These proteins are difficult to observe by definition, and it is not surprising that few have been found in studies of the stability of cellular proteins. It is possible that the degradation of such proteins contributes a substantial fraction of the peptide bonds that are rapidly turned over in growing cells.
It is now clear that the products of many of the genes in the heat shock regulon play important roles in proteolytic processes. Indeed, it is probably reasonable to view the proteolytic apparatus and the proteins of the heat shock regulon as parts of a single system responsible for ridding the cell of potentially deleterious, misfolded proteins. Proteins that find themselves in a nonnative conformation (as a result of misfolding during synthesis or of unfolding in response to stress) may either refold, aggregate, or be degraded. Heat shock proteins catalyze refolding, prevent aggregation, and, along with at least part of the proteolytic apparatus, participate in the degradation of unsalvageable proteins. Heat shock proteins probably play important roles in the cell under all conditions but especially after exposure to high temperature or to the other stresses that induce the heat shock response.
Three kinds of observations demonstrate the links between the heat shock regulon and the proteolytic apparatus. (i) The presence in the cell of abnormal proteins that are specifically recognized and degraded by the proteolytic apparatus appears to be the signal that leads to induction of the heat shock response. Accumulation of canavanine-containing proteins, of puromycin-terminated incomplete proteins, or of proteins containing streptomycin-induced translational errors leads to the induction of the heat shock response, and this induction is dependent on htpR, the gene encoding σ 32 (69). Strains accumulating secretory protein precursors show a similar elevation in the levels of heat shock proteins (251). Even the expression of a single hyperdegradable protein leads to induction of the heat shock response (69, 181). Although the inducing proteins are substrates for proteolysis, it is the presence of the protein, not its degradation, that is required for heat shock induction (181). (ii) The genes for some proteases are directly regulated as part of the regulon. As noted above, Lon, ClpP, ClpX, and OpdA are all σ 32-dependent heat shock proteins. There is also evidence that the level of another ATP-dependent protease is elevated in mutants lacking the heat shock protein encoded by dnaK (124). (iii) The levels of heat shock proteins affect the rate of protein degradation. Mutational loss of any of several heat shock proteins (GroE, DnaJ, DnaK, or GrpE) leads to a diminished rate of degradation of puromycin-induced protein fragments and canavanine-containing proteins (117, 220). Increased levels of heat shock proteins can increase the rate of degradation of puromycyl fragments, suggesting that the availability of heat shock proteins might be rate limiting for some degradation processes (220). Heat shock proteins are also involved in the degradation of normally unstable proteins. The rapid in vivo degradation of σ 32 requires DnaK (131a). Some proteins are actually degraded more rapidly in strains defective in heat shock proteins. A missense mutant LacI gene product is destabilized by the loss of DnaK (117). Although the basis for these observations is not firmly established, it is likely that they can all be explained by the ability of heat shock proteins to alter the folding or the aggregation state of other proteins. Interaction with a heat shock protein might maintain or increase the sensitivity to proteolysis of some substrates by maintaining them in a susceptible conformation or by preventing them from aggregating (puromycyl fragments, canavanine proteins, and σ 32). Other unstable proteins might interact with heat shock proteins in such a way that they are refolded into a conformation relatively resistant to protease attack (missense LacI protein).
What determines the susceptibility of a potential substrate to proteolysis? This is perhaps the key question in understanding in vivo proteolysis. Bacteria appear to have no general "marking" reaction comparable to that provided by the ubiquitin system in eukaryotic cells, and there is no evidence that substrates for proteolysis are sequestered in any way. It appears, therefore, that the specificity of recognition lies in the interaction between proteases and their substrates or, perhaps, between proteases and substrates bound to heat shock proteins. In only a few cases, however, has it been possible to demonstrate the specific degradation of physiological substrates in vitro by using purified proteins (137, 253).
Although it is not possible to specify the nature of the recognition process, it is clear that there is more than one recognition signal. As noted above, studies with mutants lacking a single protease indicate that each protease is relatively specific for a particular group of proteins. This specificity is dramatically illustrated in the phage λ system (77). λN is degraded by Lon, λO is degraded by ClpXP (but not ClpAP), cII is degraded by Hfl, cI undergoes RecA-stimulated autodegradation, and Xis is degraded by an unknown protease. Thus, substrate proteins appear to contain several different signals that target their degradation by specific proteases. Because both lon and clp mutants show diminished rates of degradation of abnormal proteins, subsets of these proteins must display these signals also, allowing them to be targeted for degradation by one of these enzymes. Since the loss of both Lon and ClpP has little effect on starvation degradation (138), other signals and other proteases must be primarily responsible for degradation under these conditions.
Clues about the nature of the signals that target proteins for proteolysis have come from studying the effects of alterations in protein structure on degradation rates. Several general classes of alterations have long been known to increase the degradation rate of normally stable proteins. These alterations include amino acid substitutions (caused by missense mutations, incorporation of amino acid analogs, translational miscoding, or posttranslational modification) and truncation (caused by nonsense mutations) or extension (caused by frameshift mutations or generated in vitro by gene fusion) of the polypeptide chain. Presumably, the common result of all of these alterations is the exposure of structural elements that are not accessible in the normal protein. Even normally stable native proteins may display such targeting signals. Many proteins normally exist in the cell in association with other proteins. In general, the production of such proteins is regulated to ensure the balanced production of all the components of the complex. When these regulatory constraints are lost (by overproduction of one component from a plasmid, for example), the imbalance created is frequently alleviated by rapid degradation of the component produced in excess. Here, too, this degradation is presumably targeted by the display of structural elements that are hidden in the complex but exposed when the protein is not associated with its normal partners.
Recent studies have probed the effects of specific changes in structure on degradation rate in greater detail. These studies have shown that quite small changes can have very large effects on degradation rates. Amino acid substitutions that replace a single Pro residue (Pro-78) in the λ repressor protein produce variants that are so rapidly degraded in the cell that repressor function is lost (188). One of these variants has been shown to be essentially wild type for dimerization and DNA binding so that the loss of in vivo function is a direct result of the degradation rate of the mutant protein. Such apparently minor structural differences appear to be responsible for surprising differences in the stability of closely related foreign proteins expressed in E. coli. Although human and rat 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase share 96% identical amino acid sequences, the human enzyme is so unstable to proteolysis in E. coli that it cannot be detected, whereas the rat enzyme is stable (127). An alteration that converts a single amino acid in the human protein to that found at the corresponding position in the rat enzyme produces a stable protein that can be expressed at high levels.
The terminal regions of the polypeptide chain appear to play a special role in determining susceptibility to degradation. In some cases, the presence of a hydrophilic C terminus seems to confer stability while a hydrophobic C-terminal sequence leads to instability (19). A protease that degrades in vitro a protein with a degradation-prone C terminus has been purified (205) (see the section on Prc, above), but mutations in the gene encoding this protease have no effect on in vivo degradation (206). There is impressive evidence that the identity of the N-terminal amino acid can profoundly affect the degradation rate (235). A series of β-galactosidase variants, each differing from the wild type only in the N-terminal amino acid, can be expressed in E. coli. All of these variants are stable to proteolytic degradation except those carrying N-terminal Phe, Leu, Trp, Tyr, Arg, or Lys. The instability of proteins with the N-terminal Arg or Lys depends on the posttranslational addition of an N-terminal Leu residue catalyzed by the leucyl, phenylalanyl-tRNA-protein transferase encoded by the aat gene (204). All of these variant proteins are stable in mutants lacking the ClpAP protease (235), so it is this enzyme which specifically recognizes these N-end rule substrates. N-end rule targeting seems to be a general pathway for proteolysis in eukaryotic microorganisms and in animal cells (74), and the N-terminal amino acids that target for proteolysis in E. coli seem to be a subset of those that confer instability in eukaryotic cells. No nonmutant E. coli proteins that are degraded via the N-end rule pathway have been identified, and, since ClpAP is dispensable for growth, the physiological significance of this pathway is uncertain.
Studies such as these illustrate the concept that amino acids important for "function" may in fact be required for proteolytic stability. It is important to recognize that one aspect of the function of every cellular protein is to have an appropriate rate of degradation. A related and perhaps equally underappreciated notion is that protein evolution is constrained by the proteolytic apparatus. A mutation that produces a protein with a usefully modified function does the cell no good if the mutant protein is degraded too rapidly to carry out this function.
Only two types of reactions account for most of the posttranslational proteolytic processing known to take place in E. coli: N-terminal Met removal and cleavage of signal peptides. In both cases, the processing reaction produces a mature product and a small, rapidly degraded peptide or a free amino acid. Outside the bacterial world, there are many examples of proteolytic processing events that generate two or more polypeptides from a single protein precursor. In E. coli, one of the few examples of such a process is the synthesis of the periplasmic protein penicillin acylase. This enzyme is encoded by the pac gene as an 846-amino-acid precursor (197). The primary gene product is processed in an ordered sequence, first by removal of a 26-amino-acid signal peptide during export to the periplasm and then by two successive endoproteolytic steps that generate the mature heterodimeric protein (containing a 209-amino-acid α subunit and a 557-amino-acid β subunit) and a 54-amino-acid peptide that is not part of the mature protein (207).
There has been substantial progress over the last few years in understanding the overall significance of proteolysis in the cell and the roles of individual proteases in proteolytic processes. However, very significant problems are still unsolved. At the simplest level, there is still considerable sorting out to be done. Not all of the proteases have been identified, and there is still uncertainty about the relationships between some of the activities that have been described biochemically. These will not be fully resolved until each activity is associated with a genetic locus and a sequence. Although there are already more enzymes than specific functions, more proteases are certain to be found.
The task of assigning specific functions to individual enzymes is far from complete. Many of the enzymes that participate in the degradation of specific phage proteins have not been assigned cellular functions. The physiological roles of many proteolytic enzymes that have been identified biochemically are unknown, and the assignment of physiological function on the basis of in vitro studies has frequently been misleading. Mutations affecting each of these activities will have to be obtained and their consequences will have to be characterized before convincing physiological roles for these enzymes can be assigned. This task is made more difficult by the fact that more than one protease may be able to function in a single process: overlapping specificities are common. There are also several functions without specific enzymes. The enzymes responsible for the turnover of proteins during starvation and the enzyme that degrades σ 32, for example, have not been identified. Each cellular compartment contains its own complement of proteolytic enzymes, and it is likely that there are compartment-specific proteolytic processes, many of which remain to be discovered.
Understanding the structural basis for specific protease susceptibility has become a feasible goal. The recognition that several different types of signals are recognized by specific proteases and the definition of the members of each degradation class should allow focused investigation of the molecular bases for protease-substrate recognition. Interactions between heat shock proteins and proteolytic substrates clearly play key roles in determining this susceptibility for some proteins, and these interactions are beginning to be defined.
The significance of proteolysis in the overall economy of the cell cannot yet be fully evaluated. There is evidence that a significant fraction of the peptide bonds synthesized are rapidly broken down. It will be important to understand more precisely what this fraction is and to determine the origin of the bonds that are subject to this rapid breakdown.
References
1. Abdullah, K. M., R. Y. Lo, and A. Mellors. 1991. Cloning, nucleotide sequence, and expression of the Pasteurella haemolytica A1 glycoprotease gene. J. Bacteriol. 173:5597–5603.
2. Adams, J. M. 1968. On the release of the formyl group from nascent protein. J. Mol. Biol. 33:571–589.
3. Affholter, J. A., V. A. Fried, and R. A. Roth. 1988. Human insulin-degrading enzyme shares structural and functional homologies with E. coli protease III. Science 242:1415–1418.
4. Amerik, A., V. K. Antonov, A. E. Gorbalenya, S. A. Kotova, T. V. Rotanova, and E. V. Shimbarevich. 1991. Site-directed mutagenesis of La protease. A catalytically active serine residue. FEBS Lett. 287:211–214.
5. Amerik, A., G. V. Petukhova, V. G. Grigorenko, I. P. Lykov, S. V. Yarovoi, V. M. Lipkin, and A. E. Gorbalenya. 1994. Cloning and sequence analysis of cDNA for a human homolog of eubacterial ATP-dependent Lon proteases. FEBS Lett. 340:25–28.
6. Anastasi, A., C. G. Knight, and A. J. Barrett. 1993. Characterization of the bacterial metalloendopeptidase pitrilysin by use of a continuous fluorescence assay. Biochem. J. 290:601–607.
7. Arfin, S. M., and R. A. Bradshaw. 1988. Cotranslational processing and protein turnover in eukaryotic cells. Biochemistry 27:7979–7984.
8. Bahl, H., H. Echols, D. B. Straus, D. Court, R. Crowl, and C. P. Georgopoulos. 1987. Induction of the heat shock response of E. coli through stabilization of sigma 32 by the phage lambda cIII protein. Genes Dev. 1:57–64.
9. Baird, L., B. Lipinska, S. Raina, and C. Georgopoulos. 1991. Identification of the Escherichia coli sohB gene, a multicopy suppressor of the HtrA (DegP) null phenotype. J. Bacteriol. 173:5763–5770.
10. Baneyx, F., and G. Georgiou. 1990. In vivo degradation of secreted fusion proteins by the Escherichia coli outer membrane protease OmpT. J. Bacteriol. 172:491–494.
11. Baneyx, F., and G. Georgiou. 1991. Construction and characterization of Escherichia coli strains deficient in multiple secreted proteases: protease III degrades high-molecular-weight substrates in vivo. J. Bacteriol. 173:2696–2703.
12. Banuett, F., and I. Herskowitz. 1987. Identification of polypeptides encoded by an Escherichia coli locus (hflA) that governs the lysis-lysogeny decision of bacteriophage lambda. J. Bacteriol. 169:4076–4085.
13. Banuett, F., M. A. Hoyt, L. McFarlane, H. Echols, and I. Herskowitz. 1986. hflB, a new Escherichia coli locus regulating lysogeny and the level of bacteriophage lambda cII protein. J. Mol. Biol. 187:213–224.
14. Baumler, A. J., J. G. Kusters, I. Stojiljkovic, and F. Heffron. 1994. Salmonella typhimurium loci involved in survival within macrophages. Infect. Immun. 62:1623–1630.
15. Bazan, J. F., L. H. Weaver, S. L. Roderick, R. Huber, and B. W. Matthews. 1994. Sequence and structure comparison suggest that methionine aminopeptidase, prolidase, aminopeptidase P, and creatinase share a common fold. Proc. Natl. Acad. Sci. USA 91:2473–2477.
16. Ben-Bassat, A., K. Bauer, S. Y. Chang, K. Myambo, A. Boosman, and S. Chang. 1987. Processing of the initiation methionine from proteins: properties of the Escherichia coli methionine aminopeptidase and its gene structure. J. Bacteriol. 169:751–757.
17. Bibi, E., P. Gros, and H. R. Kaback. 1993. Functional expression of mouse mdr1 in Escherichia coli. Proc. Natl. Acad. Sci. USA 90:9209–2013.
18. Blattner, F. R., V. Burland, G. Plunkett III, H. J. Sofia, and D. L. Daniels. 1993. Analysis of the Escherichia coli genome. IV. DNA sequence of the region from 89.2 to 92.8 minutes. Nucleic Acids Res. 21:5408–5417.
19. Bowie, J. U., and R. T. Sauer. 1989. Identification of C-terminal extensions that protect proteins from intracellular proteolysis. J. Biol. Chem. 264:7596–7602.
20. Bukhari, A.I., and D. Zipser. 1973. Mutants of Escherichia coli with a defect in the degradation of nonsense fragments. Nature (London) New Biol. 243:238–241.
21. Burley, S. K., P. R. David, and W. N. Lipscomb. 1991. Leucine aminopeptidase: bestatin inhibition and a model for enzyme-catalyzed peptide hydrolysis. Proc. Natl. Acad. Sci. USA 88:6916–6920.
22. Burley, S. K., P. R. David, R. M. Sweet, A. Taylor, and W. N. Lipscomb. 1992. Structure determination and refinement of bovine lens leucine aminopeptidase and its complex with bestatin. J. Mol. Biol. 224:113–140.
23. Burley, S. K., P. R. David, A. Taylor, and W. N. Lipscomb. 1990. Molecular structure of leucine aminopeptidase at 2.7-Å resolution. Proc. Natl. Acad. Sci. USA 87:6878–6882.
24. Caron, P. R., and L. Grossman. 1988. Potential role of proteolysis in the control of UvrABC incision. Nucleic Acids Res. 16:10903–10912.
25. Carter, T. H., and C. G. Miller. 1984. Aspartate-specific peptidases in Salmonella typhimurium: mutants deficient in peptidase E. J. Bacteriol. 159:453–459.
26. Cavard, D., C. Lazdunski, and S. P. Howard. 1989. The acylated precursor form of the colicin A lysis protein is a natural substrate of the DegP protease. J. Bacteriol. 171:6316–6322.
27. Cerritelli, S. M., D. Y. Shin, H. C. Chen, M. Gonzales, and R. J. Crouch. 1993. Proteolysis of Saccharomyces cerevisiae RNase H1 in E. coli. Biochimie 75:107–111.
28. Chang, S. Y., E. C. McGary, and S. Chang. 1989. Methionine aminopeptidase gene of Escherichia coli is essential for cell growth. J. Bacteriol. 171:4071–4072.
29. Charette, M. F., G. W. Henderson, and A. Markovitz. 1981. ATP hydrolysis-dependent protease activity of the lon (capR) protein of Escherichia coli K-12. Proc. Natl. Acad. Sci. USA 78:4728–4732.
30. Cheng, H. H., and H. Echols. 1987. A class of Escherichia coli proteins controlled by the hflA locus. J. Mol. Biol. 196:737–740.
31. Cheng, H. H., P. J. Muhlrad, M. A. Hoyt, and H. Echols. 1988. Cleavage of the cII protein of phage lambda by purified HflA protease: control of the switch between lysis and lysogeny. Proc. Natl. Acad. Sci. USA 85:7882–7886.
32. Cheng, Y. S., and D. Zipser. 1979. Purification and characterization of protease III from Escherichia coli. J. Biol. Chem. 254:4698–4706.
33. Chin, D. T., S. A. Goff, T. Webster, T. Smith, and A. L. Goldberg. 1988. Sequence of the lon gene in Escherichia coli. A heat-shock gene which encodes the ATP-dependent protease La. J. Biol. Chem. 263:11718–11728.
34. Cho, H., and J. E. Cronan, Jr. 1994. "Protease I" of Escherichia coli functions as a thioesterase in vivo. J. Bacteriol. 176:1793–1795.
35. Chung, C. H., and A. L. Goldberg. 1981. The product of the lon (capR) gene in Escherichia coli is the ATP-dependent protease, protease La. Proc. Natl. Acad. Sci. USA 78:4931–4935.
36. Chung, C. H., and A. L. Goldberg. 1982. DNA stimulates ATP-dependent proteolysis and protein-dependent ATPase activity of protease La from Escherichia coli. Proc. Natl. Acad. Sci. USA 79:795–799.
37. Chung, C. H., and A. L. Goldberg. 1983. Purification and characterization of protease So, a cytoplasmic serine protease in Escherichia coli. J. Bacteriol. 154:231–238.
38. Chung, C. H., H. E. Ives, S. Almeda, and A. L. Goldberg. 1983. Purification from Escherichia coli of a periplasmic protein that is a potent inhibitor of pancreatic proteases. J. Biol. Chem. 258:11032–11038.
39. Claverie-Martin, F., M. R. Diaz-Torres, and S. R. Kushner. 1987. Analysis of the regulatory region of the protease III (ptr) gene of Escherichia coli K-12. Gene 54:185–195.
40. Conlin, C. A. 1991. Characterization of the Salmonella typhimurium opdA gene, encoding oligopeptidase a: nucleotide sequence; identity with the Escherichia coli prlC gene; and its role in bacteriophage P22 development. Ph.D. thesis. Case Western Reserve University, Cleveland, Ohio.
41. Conlin, C. A., K. Haakensson, A. Liljas, and C. G. Miller. 1994. Cloning and nucleotide sequence of the cyclic AMP receptor protein-regulated Salmonella typhimurium pepE gene and crystallization of its product, an α-aspartyl dipeptidase. J. Bacteriol. 176:166–172.
42. Conlin, C. A., N. J. Trun, T. J. Silhavy, and C. G. Miller. 1992. Escherichia coli prlC encodes an endopeptidase and is homologous to the Salmonella typhimurium opdA gene. J. Bacteriol. 174:5881–5887.
43. Conlin, C. A., E. R. Vimr, and C. G. Miller. 1992. Oligopeptidase A is required for normal phage P22 development. J. Bacteriol. 174:5869–5880.
44. Dalbey, R. E. 1991. Leader peptidase. Mol. Microbiol. 5:2855–2860.
45. Dalbey, R. E., and G. von Heijne. 1992. Signal peptidases in prokaryotes and eukaryotes—a new protease family. Trends Biochem. Sci. 17:474–478. (Erratum, 18:25, 1993.)
46. Dalphin, M. E., B. A. Fane, M. O. Skidmore, and M. Hayashi. 1992. Proteolysis of bacteriophage φX174 prohead accessory protein gpB by Escherichia coli OmpT protease is not essential for phage maturation in vivo. J. Bacteriol. 174:2404–2406.
47. Damerau, K., and A. St. John. 1993. Role of Clp protease subunits in degradation of carbon starvation proteins in Escherichia coli. J. Bacteriol. 175:53–63.
48. Das, A., J. Urbanowski, H. Weissbach, J. Nestor, and C. Yanofsky. 1983. In vitro synthesis of the tryptophan operon leader peptides of Escherichia coli, Serratia marcescens, and Salmonella typhimurium. Proc. Natl. Acad. Sci. USA 80:2879–2883.
49. Derbyshire, K. M., M. Kramer, and N. D. Grindley. 1990. Role of instability in the cis action of the insertion sequence IS903 transposase. Proc. Natl. Acad. Sci. USA 87:4048–4052.
50. Deutch, C. E., and R. L. Soffer. 1978. Escherichia coli mutants defective in dipeptidyl carboxypeptidase. Proc. Natl. Acad. Sci. USA 75:5998–6001.
51. Diaz-Torres, M. R., C. C. Dykstra, F. Claverie-Martin, and S. R. Kushner. 1991. Extracellular release of protease III (ptr) by Escherichia coli K12. Can. J. Microbiol. 37:718–721.
52. Ding, L., A. B. Becker, A. Suzuki, and R. A. Roth. 1992. Comparison of the enzymatic and biochemical properties of human insulin-degrading enzyme and Escherichia coli protease III. J. Biol. Chem. 267:2414–2420.
53. ndo, F., and I. Matsuda. 1991. Molecular basis of prolidase (peptidase D) deficiency. Mol. Biol. Med. 8:117–127.
54. Erickson, B. D., Z. F. Burton, K. K. Watanabe, and R. R. Burgess. 1985. Nucleotide sequence of the rpsU-dnaG-rpoD operon from Salmonella typhimurium and a comparison of this sequence with the homologous operon of Escherichia coli. Gene 40:67–78.
55. Erickson, J. W., and C. A. Gross. 1989. Identification of the sigma E subunit of Escherichia coli RNA polymerase: a second alternate sigma factor involved in high-temperature gene expression. Genes Dev. 3:1462–1471.
56. Erpel, T., P. Hwang, C. S. Craik, R. J. Fletterick, and M. E. McGrath. 1992. Physical map location of the new Escherichia coli gene eco, encoding the serine protease inhibitor ecotin. J. Bacteriol. 174:1704.
57. Finch, P. W., A. Storey, K. E. Chapman, K. Brown, I. D. Hickson, and P. T. Emmerson. 1986. Complete nucleotide sequence of the Escherichia coli recB gene. Nucleic Acids Res. 14:8573–8582.
58. Fischer, H., and R. Glockshuber. 1993. ATP hydrolysis is not stoichiometrically linked with proteolysis in the ATP-dependent protease La from Escherichia coli. J. Biol. Chem. 268:22502–22507.
59. Fiss, E. H., W. C. J. Hollifield, and J. B. Neilands. 1979. Absence of ferric enterobactin receptor modification activity in mutants of Escherichia coli K-12 lacking protein a. Biochem. Biophys. Res. Commun. 91:29–34.
60. Fiss, E. H., P. Stanley-Samuelson, and J. B. Neilands. 1982. Properties and proteolysis of ferric enterobactin outer membrane receptor in Escherichia coli K12. Biochemistry 21:4517–4522.
61. Foglino, M., S. Gharbi, and A. Lazdunski. 1986. Nucleotide sequence of the pepN gene encoding aminopeptidase N of Escherichia coli. Gene 49:303–309.
62. Furuchi, T., K. Kashiwagi, H. Kobayashi, and K. Igarashi. 1991. Characteristics of the gene for a spermidine and putrescine transport system that maps at 15 min on the Escherichia coli chromosome. J. Biol. Chem. 266:20928–20933.
63. Gama, M. J., A. Toussaint, and M. L. Pato. 1990. Instability of bacteriophage Mu transposase and the role of host Hfl protein. Mol. Microbiol. 4:1891–1897.
64. Gentry, D. R., and R. R. Burgess. 1990. Overproduction and purification of the omega subunit of Escherichia coli RNA polymerase. Protein Expression Purif. 1:81–86.
65. Georgopoulos, C., and W. J. Welch. 1993. Role of the major heat shock proteins as molecular chaperones. Annu. Rev. Cell Biol. 9:601–634.
66. Geuskens, V., A. Mhammedi-Alaoui, L. Desmet, and A. Toussaint. 1992. Virulence in bacteriophage Mu: a case of trans-dominant proteolysis by the Escherichia coli Clp serine protease. EMBO J. 11:5121–5127.
67. Gharbi, S., A. Belaich, M. Murgier, and A. Lazdunski. 1985. Multiple controls exerted on in vivo expression of the pepN gene in Escherichia coli: studies with pepN-lacZ operon and protein fusion strains. J. Bacteriol. 163:1191–1195.
68. Goff, S. A., L. P. Casson, and A. L. Goldberg. 1984. Heat shock regulatory gene htpR influences rates of protein degradation and expression of the lon gene in Escherichia coli. Proc. Natl. Acad. Sci. USA 81:6647–6651.
69. Goff, S. A., and A. L. Goldberg. 1985. Production of abnormal proteins in E. coli stimulates transcription of lon and other heat shock genes. Cell 41:587–595.
70. Goff, S. A., and A. L. Goldberg. 1987. An increased content of protease La, the lon gene product, increases protein degradation and blocks growth in Escherichia coli. J. Biol. Chem. 262:4508–4515.
71. Goldberg, A. L. 1992. The mechanism and functions of ATP-dependent proteases in bacterial and animal cells. Eur. J. Biochem. 203:9–23.
72. Goldberg, A. L., K. H. Swamy, C. H. Chung, and F. S. Larimore. 1981. Proteases in Escherichia coli. Methods Enzymol. 680–702.
73. Goldberg, A. L., and L. Waxman. 1985. The role of ATP hydrolysis in the breakdown of proteins and peptides by protease La from Escherichia coli. J. Biol. Chem. 260:12029–12034.
74. Gonda, D. K., A. Bachmair, I. Wunning, J. W. Tobias, W. S. Lane, and A. Varshavsky. 1989. Universality and structure of the N-end rule. J. Biol. Chem. 264:16700–16712.
75. Gottesman, S. 1989. Genetics of proteolysis in Escherichia coli. Annu. Rev. Genet. 23:163–198.
76. Gottesman, S. 1990. Minimizing proteolysis in Escherichia coli: genetic solutions. Methods Enzymol. 185:119–129.
77. Gottesman, S., W. P. Clark, V. de Crecy-Lagard, and M. R. Maurizi. 1993. ClpX, an alternative subunit for the ATP-dependent Clp protease of Escherichia coli. Sequence and in vivo activities. J. Biol. Chem. 268:22618–22626.
78. Gottesman, S., W. P. Clark, and M. R. Maurizi. 1990. The ATP-dependent Clp protease of Escherichia coli. Sequence of clpA and identification of a Clp-specific substrate. J. Biol. Chem. 265:7886–7893.
79. Gottesman, S., and M. R. Maurizi. 1992. Regulation by proteolysis: energy-dependent proteases and their targets. Microbiol. Rev. 56:592–621.
80. Gottesman, S., C. Squires, E. Pichersky, M. Carrington, M. Hobbs, J. S. Mattick, B. Dalrymple, H. Kuramitsu, T. Shiroza, and T. Foster. 1990. Conservation of the regulatory subunit for the Clp ATP-dependent protease in prokaryotes and eukaryotes. Proc. Natl. Acad. Sci. USA 87:3513–3517.
81. Gottesman, S., and D. Zipser. 1978. Deg phenotype of Escherichia coli lon mutants. J. Bacteriol. 133:844–851.
82. Grodberg, J., and J. J. Dunn. 1989. Comparison of Escherichia coli K-12 outer membrane protease OmpT and Salmonella typhimurium E protein. J. Bacteriol. 171:2903–2905.
83. Grodberg, J., M. D. Lundrigan, D. L. Toledo, W. F. Mangel, and J. J. Dunn. 1988. Complete nucleotide sequence and deduced amino acid sequence of the ompT gene of Escherichia coli K-12. Nucleic Acids Res. 16:1209.
84. Haley, E. E. 1968. Purification and properties of a beta-aspartyl peptidase from Escherichia coli. J. Biol. Chem. 243:5748–5752.
85. Hamilton, S., and C. G. Miller. 1992. Cloning and nucleotide sequence of the Salmonella typhimurium dcp gene encoding dipeptidyl carboxypeptidase. J. Bacteriol. 174:1626–1630.
86. Hammarberg, B., T. Moks, M. Tally, A. Elmblad, E. Holmgren, M. Murby, B. Nilsson, S. Josephson, and M. Uhlen. 1990. Differential stability of recombinant human insulin-like growth factor II in Escherichia coli and Staphylococcus aureus. J. Biotechnol. 14:423–437.
87. Hanke, C., J. Hess, G. Schumacher, and W. Goebel. 1992. Processing by OmpT of fusion proteins carrying the HlyA transport signal during secretion by the Escherichia coli hemolysin transport system. Mol. Gen. Genet. 233:42–48.
88. Hara, H., Y. Nishimura, J. Kato, H. Suzuki, H. Nagasawa, A. Suzuki, and Y. Hirota. 1989. Genetic analyses of processing involving C-terminal cleavage in penicillin-binding protein 3 of Escherichia coli. J. Bacteriol. 171:5882–5889.
89. Hara, H., Y. Yamamoto, A. Higashitani, H. Suzuki, and Y. Nishimura. 1991. Cloning, mapping, and characterization of the Escherichia coli prc gene, which is involved in C-terminal processing of penicillin-binding protein 3. J. Bacteriol. 173:4799–4813.
90. Harding, V., A. Karim, N. Kaderbhai, A. Jones, A. Evans, and M. A. Kaderbhai. 1993. Processing of chimeric mammalian cytochrome b5 precursors in Escherichia coli: reaction specificity of signal peptidase and identification of an aminopeptidase in post-translocational processing. Biochem. J. 293:751–756.
91. Hatzfeld, M., H. Dodemont, U. Plessmann, and K. Weber. 1992. Truncation of recombinant vimentin by ompT. Identification of a short motif in the head domain necessary for assembly of type III intermediate filament proteins. FEBS Lett. 302:239–242.
92. Heiman, C., and C. G. Miller. 1978. Salmonella typhimurium mutants lacking protease II. J. Bacteriol. 135:588–594.
93. Henderson, T. A., P. M. Dombrosky, and K. D. Young. 1994. Artifactual processing of penicillin-binding proteins 7 and 1b by the OmpT protease of Escherichia coli. J. Bacteriol. 176:256–259.
94. Henrich, B., H. Backes, J. R. Klein, and R. Plapp. 1992. The promoter region of the Escherichia coli pepD gene: deletion analysis and control by phosphate concentration. Mol. Gen. Genet. 232:117–125.
95. Henrich, B., S. Becker, U. Schroeder, and R. Plapp. 1993. dcp gene of Escherichia coli: cloning, sequencing, transcript mapping, and characterization of the gene product. J. Bacteriol. 175:7290–7300.
96. Henrich, B., U. Monnerjahn, and R. Plapp. 1990. Peptidase D gene (pepD) of Escherichia coli K-12: nucleotide sequence, transcript mapping, and comparison with other peptidase genes. J. Bacteriol. 172:4641–4651.
97. Henrich, B., and R. Plapp. 1991. Locations of the genes from pepD through proA on the physical map of the Escherichia coli chromosome. J. Bacteriol. 173:7407–7408.
98. Herman, C., T. Ogura, T. Tomoyasu, S. Hiraga, Y. Akiyama, K. Ito, R. Thomas, R. D’Ari, and P. Bouloc. 1993. Cell growth and lambda phage development controlled by the same essential Escherichia coli gene, ftsH/hflB. Proc. Natl. Acad. Sci. USA 90:10861–10865.
99. Herskowitz, I., and D. Hagen. 1980. The lysis-lysogeny decision of phage λ: explicit programming and responsiveness. Annu. Rev. Genet. 14:399–445.
100. Hobbs, M., and J. S. Mattick. 1993. Common components in the assembly of type 4 fimbriae, DNA transfer systems, filamentous phage and protein-secretion apparatus: a general system for the formation of surface-associated protein complexes. Mol. Microbiol. 10:233–243.
101. Hollifield, W. C. J., E. H. Fiss, and J. B. Neilands. 1978. Modification of a ferric enterobactin receptor protein from the outer membrane of Escherichia coli. Biochem. Biophys. Res. Commun. 83:739–746.
102. Hoyt, M. A., D. M. Knight, A. Das, H. I. Miller, and H. Echols. 1982. Control of phage lambda development by stability and synthesis of cII protein: role of the viral cIII and host hflA, himA and himD genes. Cell 31:565–573.
103. Hussain, M., S. Ichihara, and S. Mizushima. 1982. Mechanism of signal peptide cleavage in the biosynthesis of the major lipoprotein of the Escherichia coli outer membrane. J. Biol. Chem. 257:5177–5182.
104. Hwang, B. J., K. M. Woo, A. L. Goldberg, and C. H. Chung. 1988. Protease Ti, a new ATP-dependent protease in Escherichia coli, contains protein-activated ATPase and proteolytic functions in distinct subunits. J. Biol. Chem. 263:8727–8734.
105. Ichihara, S., N. Beppu, and S. Mizushima. 1984. Protease IV, a cytoplasmic membrane protein of Escherichia coli, has signal peptide peptidase activity. J. Biol. Chem. 259:9853–9857.
106. Ichihara, S., Y. Matsubara, C. Kato, K. Akasaka, and S. Mizushima. 1993. Molecular cloning, sequencing, and mapping of the gene encoding protease I and characterization of proteinase and proteinase-defective Escherichia coli mutants. J. Bacteriol. 175:1032–1037.
107. Ichihara, S., T. Suzuki, M. Suzuki, and S. Mizushima. 1986. Molecular cloning and sequencing of the sppA gene and characterization of the encoded protease IV, a signal peptide peptidase, of Escherichia coli. J. Biol. Chem. 261:9405–9411.
108. Ishino, Y., H. Shinagawa, K. Makino, M. Amemura, and A. Nakata. 1987. Nucleotide sequence of the iap gene, responsible for alkaline phosphatase isozyme conversion in Escherichia coli and identification of the gene product. J. Bacteriol. 169:5429–5433.
109. Ito, K., S. Udaka, and H. Yamagata. 1992. Cloning, characterization, and inactivation of the Bacillus brevis lon gene. J. Bacteriol. 174:2281–2287.
110. Johnson, K., I. Charles, G. Dougan, D. Pickard, P. O’Gaora, G. Costa, T. Ali, I. Miller, and C. Hormaeche. 1991. The role of a stress-response protein in Salmonella typhimurium virulence. Mol. Microbiol. 5:401–407.
111. Mol. Microbiol. 4:2027–2033.
112. Kanatani, A., T. Masuda, T. Shimoda, F. Misoka, X. S. Lin, T. Yoshimoto, and D. Tsuru. 1991. Protease II from Escherichia coli: sequencing and expression of the enzyme gene and characterization of the expressed enzyme. J. Biochem. (Toyko) 110:315–320.
113. Katayama, Y., S. Gottesman, J. Pumphrey, S. Rudikoff, W. P. Clark, and M. R. Maurizi. 1988. The two-component, ATP-dependent Clp protease of Escherichia coli. Purification, cloning, and mutational analysis of the ATP-binding component. J. Biol. Chem. 263:15226–15236.
114. Katayama-Fujimura, Y., S. Gottesman, and M. R. Maurizi. 1987. A multiple-component, ATP-dependent protease from Escherichia coli. J. Biol. Chem. 262:4477–4485.
115. Kato, M., T. Irisawa, M. Ohtani, and M. Muramatu. 1992. Purification and characterization of proteinase In, a trypsin-like proteinase, in Escherichia coli. Eur. J. Biochem. 210:1007–1014.
116. Kaufmann, A., Y. D. Stierhof, and U. Henning. 1994. New outer membrane-associated protease of Escherichia coli K-12. J. Bacteriol. 176:359–367.
117. Keller, J. A., and L. D. Simon. 1988. Divergent effects of a dnaK mutation on abnormal protein degradation in Escherichia coli. Mol. Microbiol. 2:31–41.
118. Kim, H., S. K. Burley, and W. N. Lipscomb. 1993. Re-refinement of the X-ray crystal structure of bovine lens leucine aminopeptidase complexed with bestatin. J. Mol. Biol. 230:722–724.
119. Kirby, J. E., J. E. Trempy, and S. Gottesman. 1994. Excision of a P4-like cryptic prophage leads to Alp protease expression in Escherichia coli. J. Bacteriol. 176:2068–2081.
120. Kirsh, M., D. R. Dembinski, P. E. Hartman, and C. G. Miller. 1978. Salmonella typhimurium peptidase active on carnosine. J. Bacteriol. 134:361–374.
121. Klein, J., B. Henrich, and R. Plapp. 1986. Cloning and expression of the pepD gene of Escherichia coli. J. Gen. Microbiol. 132:2337–2343.
122. Kowit, J. D., W. N. Choy, S. P. Champe, and A. L. Goldberg. 1976. Role and location of protease I from Escherichia coli. J. Bacteriol. 128:776–784.
123. Kroh, H. E., and L. D. Simon. 1990. The ClpP component of Clp protease is the sigma 32-dependent heat shock protein F21.5. J. Bacteriol. 172:6026–6034.
124. Kroh, H. E., and L. D. Simon. 1991. Increased ATP-dependent proteolytic activity in lon-deficient Escherichia coli strains lacking the DnaK protein. J. Bacteriol. 173:2691–2695.
125. Kumagai, H., S. Nohara, H. Suzuki, W. Hashimoto, K. Yamamoto, H. Sakai, K. Sakabe, K. Fukuyama, and N. Sakabe. 1993. Crystallization and preliminary X-ray analysis of gamma-glutamyltranspeptidase from Escherichia coli K-12. J. Mol. Biol. 234:1259–1262.
126. Kuo, W. L., B. D. Gehm, and M. R. Rosner. 1990. Cloning and expression of the cDNA for a Drosophila insulin-degrading enzyme. Mol. Endocrinol. 4:1580–1591.
127. Lange, A. J., L. Li, A. M. Vargas, and S. J. Pilkis. 1993. Expression of human liver 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase in Escherichia coli. Role of N-2 proline in degradation of the protein. J. Biol. Chem. 268:8078–8084.
128. Lassen, S. F., K. K. Mortensen, and H. U. Sperling-Petersen. 1992. OmpT proteolysis of E. coli initiation factor IF2. Elimination of a cleavage site by site-directed mutagenesis. Biochem. Int. 27:601–611.
129. Lee, C. S., J. K. Hahm, B. J. Hwang, K. C. Park, D. B. Ha, S. D. Park, and C. H. Chung. 1990. Processing of Ada protein by two serine endoproteases Do and So from Escherichia coli. FEBS Lett. 262:310–312.
130. Lee, H. R., J. H. Seo, O. M. Kim, C. S. Lee, S. W. Suh, Y. M. Hong, K. Tanaka, A. Ichihara, D. B. Ha, and C. H. Chung. 1991. Molecular cloning of the ecotin gene in Escherichia coli. FEBS Lett. 287:53–56. (Erratum, 296:235, 1992.)
131. Lee, Y. S., S. C. Park, A. L. Goldberg, and C. H. Chung. 1988. Protease So from Escherichia coli preferentially degrades oxidatively damaged glutamine synthetase. J. Biol. Chem. 263:6643–6646.
131a. Liberek, K., T. P. Galitski, M. Zylicz, and C. Georgopoulos. 1992. The DnaK chaperone modulates the heat shock response of Escherichia coli by binding to the sigma-32 transcription factor. Proc. Natl. Acad. Sci. USA 89:3516–3520.
132. Lipinska, B., O. Fayet, L. Baird, and C. Georgopoulos. 1989. Identification, characterization, and mapping of the Escherichia coli htrA gene, whose product is essential for bacterial growth only at elevated temperatures. J. Bacteriol. 171:1574–1584.
133. Lipinska, B., S. Sharma, and C. Georgopoulos. 1988. Sequence analysis and regulation of the htrA gene of Escherichia coli: a sigma 32-independent mechanism of heat-inducible transcription. Nucleic Acids Res. 16:10053–10067.
134. Lipinska, B., M. Zylicz, and C. Georgopoulos. 1990. The HtrA (DegP) protein, essential for Escherichia coli survival at high temperatures, is an endopeptidase. J. Bacteriol. 172:1791–1797.
135. Lombardo, M. J., C. G. Miller, and K. E. Rudd. 1993. Physical mapping of the Escherichia coli pepT and potABCD genes. J. Bacteriol. 175:7745–7746.
136. Lowe, P. N., M. J. Page, S. Bradley, S. Rhodes, M. Sydenham, H. Paterson, and R. H. Skinner. 1991. Characterization of recombinant human Kirsten-ras (4B) p21 produced at high levels in Escherichia coli and insect baculovirus expression systems. J. Biol. Chem. 266:1672–1678.
137. Maurizi, M. R. 1987. Degradation in vitro of bacteriophage lambda N protein by Lon protease from Escherichia coli. J. Biol. Chem. 262:2696–2703.
138. Maurizi, M. R. 1992. Proteases and protein degradation in Escherichia coli. Experientia 48:178–201.
139. Maurizi, M. R., W. P. Clark, Y. Katayama, S. Rudikoff, J. Pumphrey, B. Bowers, and S. Gottesman. 1990. Sequence and structure of Clp P, the proteolytic component of the ATP-dependent Clp protease of Escherichia coli. J. Biol. Chem. 265:12536–12545.
140. Maurizi, M. R., W. P. Clark, S. H. Kim, and S. Gottesman. 1990. Clp P represents a unique family of serine proteases. J. Biol. Chem. 265:12546–12552.
141. Maurizi, M. R., P. Trisler, and S. Gottesman. 1985. Insertional mutagenesis of the lon gene in Escherichia coli: lon is dispensable. J. Bacteriol. 164:1124–1135.
142. Mazel, D., S. Pochet, and P. Marliere. 1994. Genetic characterization of polypeptide deformylase, a distinctive enzyme of eubacterial translation. EMBO J. 13:914–923.
143. McCaman, M. T., and J. D. Gabe. 1986. The nucleotide sequence of the pepN gene and its over-expression in Escherichia coli. Gene 48:145–153.
144. McCulloch, R., M. E. Burke, and D. J. Sheratt. 1994. Peptidase activity of Escherichia coli aminopeptidase A is not required for its role in Xer site-specific recombination. Mol. Microbiol. 12:241–251.
145. McDonough, K. A., and S. Falkow. 1989. A Yersinia pestis-specific DNA fragment encodes temperature-dependent coagulase and fibrinolysin-associated phenotypes. Mol. Microbiol. 3:767–775.
146. McGrath, M. E., W. M. Hines, J. A. Sakanari, R. J. Fletterick, and C. S. Craik. 1991. The sequence and reactive site of ecotin. A general inhibitor of pancreatic serine proteases from Escherichia coli. J. Biol. Chem. 266:6620–6625.
147. McHugh, G. L., and C. G. Miller. 1974. Isolation and characterization of proline peptidase mutants of Salmonella typhimurium. J. Bacteriol. 120:364–371.
148. Menon, A. S., and A. L. Goldberg. 1987. Protein substrates activate the ATP-dependent protease La by promoting nucleotide binding and release of bound ADP. J. Biol. Chem. 262:14929–14934.
149. Menon, A. S., L. Waxman, and A. L. Goldberg. 1987. The energy utilized in protein breakdown by the ATP-dependent protease (La) from Escherichia coli. J. Biol. Chem. 262:722–726.
150. Mierau, I., A. J. Haandrikman, O. Velterop, P. S. Tan, K. L. Leenhouts, W. N. Konigs, G. Venema, and J. Kok. 1994. Tripeptidase gene (pepT) of Lactococcus lactis: molecular cloning and nucleotide sequencing of pepT and construction of a chromosomal deletion mutant. J. Bacteriol. 176:2854–2861.
151. Miller, C. G. 1987. Protein degradation and proteolytic modification, p. 680–691. In F. C. Neidhardt, J. L. Ingraham, K. B. Low, B. Magasanik, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology. American Society for Microbiology, Washington, D.C.
151a. Miller, C. G., and C. A. Conlin. 1994. Signal peptide hydrolases. In G. von Heijne (ed.) Signal Peptidases. R. G. Landes, Austin, Tex.
152. Miller, C. G., and L. Green. 1983. Degradation of proline peptides in peptidase-deficient strains of Salmonella typhimurium. J. Bacteriol. 153:350–356.
153. Miller, C. G., C. Heiman, and C. Yen. 1976. Mutants of Salmonella typhimurium deficient in an endoprotease. J. Bacteriol. 127:490–497.
154. Miller, C. G., A. M. Kukral, J. L. Miller, and N. R. Movva. 1989. pepM is an essential gene in Salmonella typhimurium. J. Bacteriol. 171:5215–5217.
155. Miller, C. G., and K. Mackinnon. 1974. Peptidase mutants of Salmonella typhimurium. J. Bacteriol. 120:355–363.
156. Miller, C. G., J. L. Miller, and D. A. Bagga. 1991. Cloning and nucleotide sequence of the anaerobically regulated pepT gene of Salmonella typhimurium. J. Bacteriol. 173:3554–3558.
157. Miller, C. G., K. L. Strauch, A. M. Kukral, J. L. Miller, P. T. Wingfield, G. J. Mazzei, R. C. Werlen, P. Graber, and N. R. Movva. 1987. N-terminal methionine-specific peptidase in Salmonella typhimurium. Proc. Natl. Acad. Sci. USA 84:2718–2722.
158. Minton, N. P., T. Atkinson, C. J. Bruton, and R. F. Sherwood. 1984. The complete nucleotide sequence of the Pseudomonas gene coding for carboxypeptidase G2. Gene 31:31–38. (Erratum, 42:353, 1986.)
159. Misra, R., A. Peterson, T. Ferenci, and T. J. Silhavy. 1991. A genetic approach for analyzing the pathway of LamB assembly into the outer membrane of Escherichia coli. J. Biol. Chem. 266:13592–13597.
160. Mizusawa, S., and S. Gottesman. 1983. Protein degradation in Escherichia coli: the lon gene controls the stability of SulA protein. Proc. Natl. Acad. Sci. USA 80:358–362.
161. Movva, N. R., D. Semon, C. Meyer, E. Kawashima, P. Wingfield, J. L. Miller, and C. G. Miller. 1990. Cloning and nucleotide sequence of the Salmonella typhimurium pepM gene. Mol. Gen. Genet. 223:345–348.
162. Munoa, F. J., K. W. Miller, R. Beers, M. Graham, and H. C. Wu. 1991. Membrane topology of Escherichia coli prolipoprotein signal peptidase (signal peptidase II). J. Biol. Chem. 266:17667–17672.
163. Nagasawa, H., Y. Sakagami, A. Suzuki, H. Suzuki, H. Hara, and Y. Hirota. 1989. Determination of the cleavage site involved in C-terminal processing of penicillin-binding protein 3 of Escherichia coli. J. Bacteriol. 171:5890–5893.
164. Nakahigashi, K., and H. Inokuchi. 1990. Nucleotide sequence between the fadB gene and the rrnA operon from Escherichia coli. Nucleic Acids Res. 18:6439.
165. Nakata, A., H. Shinagawa, and F. G. Rothman. 1987. Molecular mechanism of isozyme formation of alkaline phosphatase in Escherichia coli, p. 139–141. In A. Torriani-Gorini, F. G. Rothman, S. Silver, A. Wright, and E. Yagil (ed.), Phosphate Metabolism and Cellular Regulation in Microorganisms. American Society for Microbiology, Washington, D.C.
166. Nakata, N., T. Tobe, I. Fukuda, T. Suzuki, K. Komatsu, M. Yoshikawa, and C. Sasakawa. 1993. The absence of a surface protease, OmpT, determines the intercellular spreading ability of Shigella: the relationship between the ompT and kcpA loci. Mol. Microbiol. 9:459–468.
167. Nesin, M., J. R. Lupski, P. Svec, and G. N. Godson. 1987. Possible new genes as revealed by molecular analysis of a 5-kb Escherichia coli chromosomal region 5' to the rpsU-dnaG-rpoD macromolecular-synthesis operon. Gene 51:149–161.
168. Noble, J. A., M. A. Innis, E. V. Koonin, K. E. Rudd, F. Banuett, and I. Herskowitz. 1993. The Escherichia coli hflA locus encodes a putative GTP-binding protein and two membrane proteins, one of which contains a protease-like domain. Proc. Natl. Acad. Sci. USA 90:10866–10870.
169. Novak, P., and I. K. Dev. 1988. Degradation of a signal peptide by protease IV and oligopeptidase A. J. Bacteriol. 170:5067–5075.
170. Novak, P., P. H. Ray, and I. K. Dev. 1986. Localization and purification of two enzymes from Escherichia coli capable of hydrolyzing a signal peptide. J. Biol. Chem. 261:420–427.
171. Pacaud, M. 1976. Purification of protease II from Escherichia coli by affinity chromatography and separation of two enzyme species from cells harvested at late log phase. Eur. J. Biochem. 64:199–204.
172. Pacaud, M. 1982. Identification and localization of two membrane-bound esterases from Escherichia coli. J. Bacteriol. 149:6–14.
173. Pacaud, M. 1982. Purification and characterization of two novel proteolytic enzymes in membranes of Escherichia coli. Protease IV and protease V. J. Biol. Chem. 257:4333–4339.
174. Pacaud, M., and C. Richaud. 1975. Protease II from Escherichia coli. Purification and characterization. J. Biol. Chem. 250:7771–7779.
175. Pacaud, M., S. Sibilli, and G. Bras. 1976. Protease I from Escherichia coli. Some physicochemical properties and substrate specificity. Eur. J. Biochem. 69:141–151.
176. acaud, M., and J. Uriel. 1971. Isolation and some properties of a proteolytic enzyme from Escherichia coli (protease I). Eur. J. Biochem. 23:435–442.
177. Paces, V., L. E. Rosenberg, W. A. Fenton, and F. Kalousek. 1993. The beta subunit of the mitochondrial processing peptidase from rat liver: cloning and sequencing of a cDNA and comparison with a proposed family of metallopeptidases. Proc. Natl. Acad. Sci. USA 90:5355–5358.
178. Palmer, S. M., and A. C. St. John. 1987. Characterization of a membrane-associated serine protease in Escherichia coli. J. Bacteriol. 169:1474–1479.
179. Park, J. H., Y. S. Lee, C. H. Chung, and A. L. Goldberg. 1988. Purification and characterization of protease Re, a cytoplasmic endoprotease in Escherichia coli. J. Bacteriol. 170:921–926.
180. Parsell, D. A., Y. Sanchez, J. D. Stitzel, and S. Lindquist. 1991. Hsp104 is a highly conserved protein with two essential nucleotide-binding sites. Nature (London) 353:270–273.
181. Parsell, D. A., and R. T. Sauer. 1989. Induction of a heat shock-like response by unfolded protein in Escherichia coli: dependence on protein level not protein degradation. Genes Dev. 3:1226–1232.
182. Parsell, D. A., K. R. Silber, and R. T. Sauer. 1990. Carboxy-terminal determinants of intracellular protein degradation. Genes Dev. 4:277–286.
183. Phillips, T. A., R. A. VanBogelen, and F. C. Neidhardt. 1984. lon gene product of Escherichia coli is a heat shock protein. J. Bacteriol. 159:283–287.
184. Polgar, L., A. Szigetvari, M. Low, I. Korodi, and E. Balla. 1991. Metalloendopeptidase QG. Isolation from Escherichia coli and characterization. Biochem. J. 273:725–731.
185. Postle, K. 1990. TonB and the gram-negative dilemma. Mol. Microbiol. 4:2019–2025.
186. Rawlings, N. D., and A. J. Barrett. 1993. Evolutionary families of peptidases. Biochem. J. 290:205–218.
187. Regnier, P. 1981. The purification of protease IV of E. coli and the demonstration that it is an endoproteolytic enzyme. Biochem. Biophys. Res. Commun. 99:1369–1376.
188. Reidhaar-Olson, J. F., D. A. Parsell, and R. T. Sauer. 1990. An essential proline in lambda repressor is required for resistance to intracellular proteolysis. Biochemistry 29:7563–7571.
189. Richardson, D. J., A. Aoyama, and M. Hayashi. 1988. Proteolysis of bacteriophage φX174 prohead protein gpB by a protease located in the Escherichia coli outer membrane. J. Bacteriol. 170:5564–5571.
190. Roderick, S. L., and B. W. Matthews. 1993. Structure of the cobalt-dependent methionine aminopeptidase from Escherichia coli: a new type of proteolytic enzyme. Biochemistry 32:3907–3912.
191. Roseman, J. E., and R. L. Levine. 1987. Purification of a protease from Escherichia coli with specificity for oxidized glutamine synthetase. J. Biol. Chem. 262:2101–2110.
192. Rupprecht, K. R., G. Gordon, M. Lundrigan, R. C. Gayda, A. Markovitz, and C. Earhart. 1983. ompT: Escherichia coli K-12 structural gene for protein a(3b). J. Bacteriol. 153:1104–1106.
193. Samson, L. 1992. The suicidal DNA repair methyltransferases of microbes. Mol. Microbiol. 6:825–831.
194. Sarimo, S. S., and M. J. Pine. 1969. Taxonomic comparison of the amino termini of microbial cell proteins. J. Bacteriol. 98:368–374.
195. Sauer, R. T., K. Hehir, R. S. Stearman, M. A. Weiss, A. Jeitler-Nilsson, E. G. Suchanek, and C. O. Pabo. 1986. An engineered intersubunit disulfide enhances the stability and DNA binding of the N-terminal domain of lambda repressor. Biochemistry 25:5992–5998.
196. Schoulaker-Schwarz, R., L. Dekel-Gorodetsky, and K. H. Engelberg. 1991. An additional function for bacteriophage lambda rex: the rexB product prevents degradation of the lambda O protein. Proc. Natl. Acad. Sci. USA 88:4996–4500.
197. Schumacher, G., D. Sizmann, H. Haug, P. Buckel, and A. Bock. 1986. Penicillin acylase from E. coli: unique gene-protein relation. Nucleic Acids Res. 14:5713–5727.
198. Sedgwick, B. 1989. In vitro proteolytic cleavage of the Escherichia coli Ada protein by the ompT gene product. J. Bacteriol. 171:2249–2251.
199. Seoane, A., A. Sabbaj, L. M. McMurry, and S. B. Levy. 1992. Multiple antibiotic susceptibility associated with inactivation of the prc gene. J. Bacteriol. 174:7844–7847.
200. Seol, J. H., S. K. Woo, E. M. Jung, S. J. Yoo, C. S. Lee, K. J. Kim, K. Tanaka, A. Ichihara, D. B. Ha, and C. H. Chung. 1991. Protease Do is essential for survival of Escherichia coli at high temperatures: its identity with the htrA gene product. Biochem. Biophys. Res. Commun. 176:730–736.
201. Seymour, J. L., R. N. Lindquist, M. S. Dennis, B. Moffat, D. Yansura, D. Reilly, M. E. Wessinger, and R. A. Lazarus. 1994. Ecotin is a potent anticoagulant and reversible tight-binding inhibitor of factor Xa. Biochemistry 33:3949–3958.
202. Shapiro, J. A. 1993. A role for the Clp protease in activating Mu-mediated DNA rearrangements. J. Bacteriol. 175:2625–2631.
203. Sherman, F., J. W. Stewart, and S. Tsunasawa. 1985. Methionine or not methionine at the beginning of a protein. Bioessays 3:27–31.
204. Shrader, T. E., J. W. Tobias, and A. Varshavsky. 1993. The N-end rule in Escherichia coli: cloning and analysis of the leucyl, phenylalanyl-tRNA-protein transferase gene aat. J. Bacteriol. 175:4364–4374.
205. Silber, K. R., K. C. Keiler, and R. T. Sauer. 1992. Tsp: a tail-specific protease that selectively degrades proteins with nonpolar C termini. Proc. Natl. Acad. Sci. USA 89:295–299.
206. Silber, K. R., and R. T. Sauer. 1994. Deletion of the prc (tsp) gene provides evidence for additional tail-specific proteolytic activity in Escherichia coli K-12. Mol. Gen. Genet. 242:237–240.
207. Sizmann, D., C. Keilmann, and A. Bock. 1990. Primary structure requirements for the maturation in vivo of penicillin acylase from Escherichia coli ATCC 11105. Eur. J. Biochem. 192:143–151.
208. Skare, J. T., B. M. Ahmer, C. L. Seachord, R. P. Darveau, and K. Postle. 1993. Energy transduction between membranes. TonB, a cytoplasmic membrane protein, can be chemically cross-linked in vivo to the outer membrane receptor FepA. J. Biol. Chem. 268:16302–16308.
209. Skare, J. T., and K. Postle. 1991. Evidence for a TonB-dependent energy transduction complex in Escherichia coli. Mol. Microbiol. 5:2883–2890.
210. Skorupski, K., J. Tomaschewski, W. Ruger, and L. D. Simon. 1988. A bacteriophage T4 gene which functions to inhibit Escherichia coli Lon protease. J. Bacteriol. 170:3016–3024.
211. Sodeinde, O. A., and J. D. Goguen. 1989. Nucleotide sequence of the plasminogen activator gene of Yersinia pestis: relationship to ompT of Escherichia coli and gene E of Salmonella typhimurium. Infect. Immun. 57:1517–1523.
212. Spiro, S., and J. R. Guest. 1990. FNR and its role in oxygen-regulated gene expression in Escherichia coli. FEMS Microbiol. Rev. 6:399–428.
213. Squires, C., and C. L. Squires. 1992. The Clp proteins: proteolysis regulators or molecular chaperones? J. Bacteriol. 174:1081–1085.
214. Stadtman, E. R. 1990. Covalent modification reactions are marking steps in protein turnover. Biochemistry 29:6323–6331.
215. Stirling, C. J., S. D. Colloms, J. F. Collins, G. Szatmari, and D. J. Sherratt. 1989. xerB, an Escherichia coli gene required for plasmid ColE1 site-specific recombination, is identical to pepA, encoding aminopeptidase A, a protein with substantial similarity to bovine lens leucine aminopeptidase. EMBO J. 8:1623–1627.
216. Strauch, K. L., and J. Beckwith. 1988. An Escherichia coli mutation preventing degradation of abnormal periplasmic proteins. Proc. Natl. Acad. Sci. USA 85:1576–1580.
217. Strauch, K. L., T. H. Carter, and C. G. Miller. 1983. Overproduction of Salmonella typhimurium peptidase T. J. Bacteriol. 156:743–751.
218. Strauch, K. L., K. Johnson, and J. Beckwith. 1989. Characterization of degP, a gene required for proteolysis in the cell envelope and essential for growth of Escherichia coli at high temperature. J. Bacteriol. 171:2689–2696.
219. Strauch, K. L., and C. G. Miller. 1983. Isolation and characterization of Salmonella typhimurium mutants lacking a tripeptidase (peptidase T). J. Bacteriol. 154:763–771.
220. Straus, D. B., W. A. Walter, and C. A. Gross. 1988. Escherichia coli heat shock gene mutants are defective in proteolysis. Genes Dev. 2:1851–1858.
221. Strom, M. S., D. N. Nunn, and S. Lory. 1993. A single bifunctional enzyme, PilD, catalyzes cleavage and N-methylation of proteins belonging to the type IV pilin family. Proc. Natl. Acad. Sci. USA 90:2404–2408.
222. Sugimura, K., and N. Higashi. 1988. A novel outer-membrane-associated protease in Escherichia coli. J. Bacteriol. 170:3650–3654.
223. Sugimura, K., and T. Nishihara. 1988. Purification, characterization, and primary structure of Escherichia coli protease VII with specificity for paired basic residues: identity of protease VII and OmpT. J. Bacteriol. 170:5625–5632.
224. Suzuki, C. K., K. Suda, N. Wang, and G. Schatz. 1994. Requirement for the yeast gene LON in intramitochondrial proteolysis and maintenance of respiration. Science 264:891.
225. Suzuki, H., W. Hashimoto, and H. Kumagai. 1993. Escherichia coli K-12 can utilize an exogenous γ-glutamyl peptide as an amino acid source, for which γ-glutamyltranspeptidase is essential. J. Bacteriol. 175:6038–6040.
226. Suzuki, H., H. Kumagai, T. Echigo, and T. Tochikura. 1988. Molecular cloning of Escherichia coli K-12 ggt and rapid isolation of gamma-glutamyltranspeptidase. Biochem. Biophys. Res. Commun. 150:33–38.
227. Suzuki, H., H. Kumagai, T. Echigo, and T. Tochikura. 1989. DNA sequence of the Escherichia coli K-12 gamma-glutamyltranspeptidase gene, ggt. J. Bacteriol. 171:5169–5172.
228. Suzuki, H., H. Kumagai, and T. Tochikura. 1986. γ-Glutamyltranspeptidase from Escherichia coli K-12: formation and localization. J. Bacteriol. 168:1332–1335.
229. Suzuki, H., H. Kumagai, and T. Tochikura. 1987. Isolation, genetic mapping, and characterization of Escherichia coli K-12 mutants lacking γ-glutamyltranspeptidase. J. Bacteriol. 169:3926–3931.
230. Suzuki, T., A. Itoh, S. Ichihara, and S. Mizushima. 1987. Characterization of the sppA gene coding for protease IV, a signal peptide peptidase of Escherichia coli. J. Bacteriol. 169:2523–2528.
231. Swamy, K. H., C. H. Chung, and A. L. Goldberg. 1983. Isolation and characterization of protease do from Escherichia coli, a large serine protease containing multiple subunits. Arch. Biochem. Biophys. 224:543–554.
232. Swamy, K. H., and A. L. Goldberg. 1981. E. coli contains eight soluble proteolytic activities, one being ATP dependent. Nature (London) 292:652–654.
233. Swamy, K. H., and A. L. Goldberg. 1982. Subcellular distribution of various proteases in Escherichia coli. J. Bacteriol. 149:1027–1033.
234. Tanoue, A., F. Endo, and I. Matsuda. 1990. Structural organization of the gene for human prolidase (peptidase D) and demonstration of a partial gene deletion in a patient with prolidase deficiency. J. Biol. Chem. 265:11306–11311.
235. Tobias, J. W., T. E. Shrader, G. Rocap, and A. Varshavsky. 1991. The N-end rule in bacteria. Science 254:1374–1377.
236. Tojo, N., S. Inouye, and T. Komano. 1993. Cloning and nucleotide sequence of the Myxococcus xanthus lon gene: indispensability of lon for vegetative growth. J. Bacteriol. 175:2271–2277.
237. Tojo, N., S. Inouye, and T. Komano. 1993. The lonD gene is homologous to the lon gene encoding an ATP-dependent protease and is essential for the development of Myxococcus xanthus. J. Bacteriol. 175:4545–4549.
238. Trempy, J. E., and S. Gottesman. 1989. Alp, a suppressor of lon protease mutants in Escherichia coli. J. Bacteriol. 171:3348–3353.
239. Trempy, J. E., J. E. Kirby, and S. Gottesman. 1994. Alp suppression of Lon: dependence on the slpA gene. J. Bacteriol. 176:2061–2067.
240. Trun, N. J., and T. J. Silhavy. 1987. Characterization and in vivo cloning of prlC, a suppressor of signal sequence mutations in Escherichia coli K12. Genetics 116:513–521.
241. Trun, N. J., and T. J. Silhavy. 1989. PrlC, a suppressor of signal sequence mutations in Escherichia coli, can direct the insertion of the signal sequence into the membrane. J. Mol. Biol. 205:665–676.
242. Tsunasawa, S., J. W. Stewart, and F. Sherman. 1985. Amino-terminal processing of mutant forms of yeast iso-1-cytochrome c. The specificities of methionine aminopeptidase and acetyltransferase. J. Biol. Chem. 260:5382–5391.
243. van Alen-Boerrigter, I. J., R. Baankreis, and W. M. de Vos. 1991. Characterization and overexpression of the Lactococcus lactis pepN gene and localization of its product, aminopeptidase N. Appl. Environ. Microbiol. 57:2555–2561.
243a. Vimr, E. R. 1981. Studies on mutants of Salmonella typhimurium deficient in two proteolytic enzymes. Ph.D. thesis. Case Western Reserve University, Cleveland, Ohio.
244. Vimr, E. R., L. Green, and C. G. Miller. 1983. Oligopeptidase-deficient mutants of Salmonella typhimurium. J. Bacteriol. 153:1259–1265.
245. Vogt, V. M. 1970. Purification and properties of an aminopeptidase from Escherichia coli. J. Biol. Chem. 245:4760–4769.
246. Wadensten, H., A. Ekebacke, B. Hammarberg, E. Holmgren, C. Kalderen, M. Tally, T. Moks, M. Uhlen, S. Josephson, and M. Hartmanis. 1991. Purification and characterization of recombinant human insulin-like growth factor II (IGF-II) expressed as a secreted fusion protein in Escherichia coli. Biotechnol. Appl. Biochem. 13:412–421.
247. Walker, G. C. 1987. The SOS response of Escherichia coli, p. 1346–1357. In F. C. Neidhardt, J. L. Ingraham, K. B. Low, B. Magasanik, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology. American Society for Microbiology, Washington, D.C.
248. Waller, J.-P. 1963. The NH2-terminal residues of the proteins from cell-free extracts of E. coli. J. Mol. Biol. 7:483–496.
249. Wang, N., S. Gottesman, M. C. Willingham, M. M. Gottesman, and M. R. Maurizi. 1993. A human mitochondrial ATP-dependent protease that is highly homologous to bacterial Lon protease. Proc. Natl. Acad. Sci. USA 90:11247–11251.
250. Whitchurch, C. B., M. Hobbs, S. P. Livingston, V. Krishnapillai, and J. S. Mattick. 1991. Characterisation of a Pseudomonas aeruginosa twitching motility gene and evidence for a specialised protein export system widespread in eubacteria. Gene 101:33–44.
251. Wild, J., W. A. Walter, C. A. Gross, and E. Altman. 1993. Accumulation of secretory protein precursors in Escherichia coli induces the heat shock response. J. Bacteriol. 175:3992–3997.
252. Wingfield, P., P. Graber, G. Turcatti, N. R. Movva, M. Pelletier, S. Craig, K. Rose, and C. G. Miller. 1989. Purification and characterization of a methionine-specific aminopeptidase from Salmonella typhimurium. Eur. J. Biochem. 180:23–32.
253. Wojtkowiak, D., C. Georgopoulos, and M. Zylicz. 1993. Isolation and characterization of ClpX, a new ATP-dependent specificity component of the Clp protease of Escherichia coli. J. Biol. Chem. 268:22609–22617.
254. Woo, K. M., W. J. Chung, D. B. Ha, A. L. Goldberg, and C. H. Chung. 1989. Protease Ti from Escherichia coli requires ATP hydrolysis for protein breakdown but not for hydrolysis of small peptides. J. Biol. Chem. 264:2088–2091.
255. Yaron, A. 1976. Dipeptidyl carboxypeptidase from Escherichia coli. Methods Enzymol. 45:599–610.
256. Yaron, A., and D. Mlynar. 1968. Aminopeptidase-P. Biochem. Biophys. Res. Commun. 32:658–663.
257. Yaron, A., D. Mylnar, and A. Berger. 1972. A dipeptidocarboxypeptidase from E. coli. Biochem. Biophys. Res. Commun. 47:897–902.
258. Yen, C., L. Green, and C. G. Miller. 1980. Degradation of intracellular protein in Salmonella typhimurium peptidase mutants. J. Mol. Biol. 143:21–33.
259. Yen, C., L. Green, and C. G. Miller. 1980. Peptide accumulation during growth of peptidase deficient mutants. J. Mol. Biol. 143:35–48.
260. Yoshimoto, T., N. Murayama, T. Honda, H. Tone, and D. Tsuru. 1988. Cloning and expression of aminopeptidase P gene from Escherichia coli HB101 and characterization of expressed enzyme. J. Biochem. (Tokyo) 104:93–97.
261. Yoshimoto, T., H. Tone, T. Honda, K. Osatomi, R. Kobayashi, and D. Tsuru. 1989. Sequencing and high expression of aminopeptidase P gene from Escherichia coli HB101. J. Biochem. (Tokyo) 105:412–416.
262. Yu, G. Q., and J. S. Hong. 1986. Identification and nucleotide sequence of the activator gene of the externally induced phosphoglycerate transport system of Salmonella typhimurium. Gene 45:51–57.
263. Zdanovsky, A. G., O. I. Kulaeva, and N. K. Yankovsky. 1992. Construction and expression of diphtheria toxin-encoding gene derivatives in Escherichia coli. Gene 116:81–86.
264. Zehnbauer, B. A., E. C. Foley, G. W. Henderson, and A. Markovitz. 1981. Identification and purification of the Lon+ (CapR+) gene product, a DNA-binding protein. Proc. Natl. Acad. Sci. USA 78:2043–2047.
265. Zhao, G. P., and R. L. Somerville. 1993. An amino acid switch (Gly281→Arg) within the "hinge" region of the tryptophan synthase beta subunit creates a novel cleavage site for the OmpT protease and selectively diminishes affinity toward a specific monoclonal antibody. J. Biol. Chem. 268:14912–14920.
266. Zorick, T. S., and H. Echols. 1991. Membrane localization of the HflA regulatory protease of Escherichia coli by immunoelectron microscopy. J. Bacteriol. 173:6307–6310.
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