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Cloning, nucleotide sequence determination, and transcriptional analysis of the histidase structural gene fromStreptomyees griscus

Wu, Pen-Chaur, Ph.D.

The Ohio State University, 1994

UMI 300 N. Zeeb Rd. Ann Arbor. M I 48106 Cloning, Nucleotide Sequence Determination, and Tranacriptlonal Analyals

of the Hiatfdaee Structural Gene from Sfrepfomycesg r lf u t

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree of Doctor of Philosophy in the Graduate

School of The Ohio State University

By

Pen-ChaurWu, B.S., M.A. *****

The Ohio State University

1994

Dissertation Committee Approved By

Charles J. Daniels

Aldis Darzins Adviser

Kathleen E. Kendrick Department of Microbiology

William R. Strohl DEDICATION

To my wife, Lily, and my parents

ii ACKNOWLEDGMENTS

I thank my advisor, Dr. Kathleen E. Kendrick, for her advice and guidance for the past five years. I also thank my committee members, Drs. Charles J. Daniels, Aldis Darzins, and

William R. Strohl, as well as Drs. Samuel J. Black and F. Robert Tabita for their comments and suggestions about my research project. I am indebted to Dr. Peter J.

White and my lab colleagues for their technical assistance and encouragement.

iii VITA

January 19, 1953 ...... Bom - Tainan, Taiwan, Republic of China

1977 ...... B, S. in Biology, National Taiwan Normal University, Taipei, Taiwan, ROC

1977 - 1979 ...... Second Lieutenant ROC Army

1979 -- 1960 ...... Teaching Staff, Sun-Sun Junior High School, Tainan County, Taiwan,

ROC

1964 ...... M.A. in Microbiology, California State University, Fresno, California

1964 ~ 1966 ...... Scientist and Microbial Assay Supervisor, Cyanamid Taiwan Corp.,

Hsinchu, Taiwan, ROC

1986 -- present...... Graduate Teaching and Research Associate, Department of

Microbiology. The Ohio State University, Columbus, Ohio

PUBLICATIONS

Wu. P.-C., T.A. Kroening, P.J. White, and K.E. Kendrick. 1992. Purification of histidase from Sfrepfomyces

griseus and nucleotide sequence of thehutH structural gene. J. Bactehol. 174:1647*1655

Wu, P.-C., T.A. Kroening, P.J. White, and K.E. Kendrick. 1992. Histidine ammonia-lyase from Streptomyces

griamua Gene 115:19-25.

FIELDS OF STUDY

Major Field: Microbiology

Studies in gene regulation in streptomycetes, Dr. Kathleen E. Kendrick

iv TABLE OF CONTENTS

DEDICATION...... ii

ACKNOWLEDGMENTS...... iii

VITA...... iv

UST OF TABLES ...... viii

LIST OF FIGURES ...... ix

CHAPTER I. INTRODUCTION...... 1

Histidine Catabolism ...... 1

Biochemical Analysis of Histidase ...... 4

Genetic Analysis of Histidine Utilization ...... 7

Biological Importance of Sfrepfomyces ...... 16

Nitrogen Metabolism in Streptomyces...... 16

CHAPTER II. MATERIALS AND METHODS...... 26

Bacterial Strains, Plasmids, and Bacteriophage ...... 26

Media and Culture Conditions ...... 29

Recombinant DNA Techniques ...... 31

Isolation of DNA ...... 31

Generation and Manipulation of DNA Fragments . 33

Hybridization Analysis ...... 38

Southern Hybridization ...... 38

v Plaque Hybridization ...... 39

Northern Hybridization ...... 42

Colony Hybridization ...... 42

Nucleotide Sequence Determination ...... 43

Bacterial Transformations ...... 44

RNA Techniques ...... 46

Run-off Transcription ...... 46

Isolation of RNA ...... 47

SI Mapping and Primer Extension Analysis 46

Preparation of Cell Extracts and Enzyme Assays ...... 50

CHAPTER III. RESULTS...... 52

Cloning the hutH Gene from $. griseu s ...... 52

Features of the Nucleotide Sequence of h u tH ...... 63

Complementation Analysis ...... 69

Repressor Titration Experiments ...... 92

Transcriptional Analysis of the hutH G en e ...... 94

Molecular Characterization of S. griseus SKK896, a Hut*

M u ta n t ...... 106

CHAPTER IV. DISCUSSION ...... 109

Summary ...... 129

LIST OF REFERENCES...... 130

APPENDICES

A. Maps of Plasmid Vectors Used in This Study ...... 140

B. List of Plasmids Constructed in This Study ...... 151

vi Isolation of Hut’ Mutants List ot Tables

Table Page

1. Bacterial strains used ...... 27

2. Plasmid vectors and bacteriophage used ...... 28

3. Media used ...... 30

4. Hut enzyme activities in transformants of SKK821 and

SKKB96 ...... 93

5. Titration of a negative regulatory factor by upstream

sequences in SKK821 and SKK896 ...... 95

6. Promoter activity of the hutH upstream sequences using

promoter-probe vectors plJ486 and plJ487 ...... 98

7. Promoter activity of S. griseus transformants containing the

hutH upstream sequences in the promoter-probe

vector pXE4 ...... 100

8. Suppression of the defect in SKK896 at high copy

number ...... 107

9. List of plasmids constructed ...... 152

10. List of Hut'Mutants of S. griseus ...... 164

11. Activities of the Hut enzymes in SKK821 and Hut' mutants

of S. g rise u s ...... 165

viii List of Flgurss

Figure Pag*

1. Pathways of L-histidine utilization in bacteria ...... 2

2. Organization and transcriptional units of the bacterial hut

g e n e s ...... 8

3. Life cycle of Streptomyces...... 17

4. Metabolism of L-histidine in streptomycetes ...... 21

5. Degenerate oligonucleotide probe used for cloning hutH 40

6. Southern analysis of total genomic DNA from S. griseus

SKK821 probed with oligonucleotides specific to the

N-terminus of hutH ...... 53

7. Identification of hutH from a library of genomic DNA from

S. griseus SKK821 ...... 55

8. Southern analysis of AKK500 and AKK501 with the

oligonucleotide probe ...... 57

9. Restriction maps of pKKSOO and pKK501 ...... 59

10. Strategy used to clone hutH from S. griseus SKKB21 . . . 61

11. Determination of the direction of transcription of hutH in

pK K 501 ...... 64

ix 12. Construction and restriction maps of pKK538 and

pKK540 ...... 66

13. Strategy for the determination of the nucleotide sequence

of the 3.45 kb fcoRI/H/ndlll fragment ...... 69

14. Nucleotide and deduced amino acid sequences of the 3.45

kb EcoRI/H/ndlll fragment ...... 71

15. Identification of protein-coding sequences in the 3.45 kb

EcoRI/H/ndlll fragment by Frame analysis 75

16. Alignment of the deduced amino acid sequence of ORF1

with ORF183 from Streptomyces ambofaciens . . . 77

17. Alignment of histidase primary structures ...... 79

18. Dendrogram of the relatedness of amino acid sequences of

histidine ammonia-fyases from four species and

phenylalanine ammonia-iyases from 8 species of

plants and fungi ...... 83

19. Features of the nucleotide sequence of the 3.45 kb

EcoRI/H/ndlll fragm ent ...... 85

20. Southern hybridization of total genomic DNA from S.

griseus SKKB21 with X DNA ...... 87

21. Southern analysis of total genomic DNA from S. griseus

and S. coeticoior with the hutH structural gene as

probe ...... 90

22. Boundaries of DNA fragments used in promoter-probe

studies ...... 96

x 23. Determination of the length of hutH m R N A ...... 102

24. Identification of the hutH transcription start site ...... 104

25. Consensus sequence of E. co/Mike streptomycete

promoters and the nucleotide sequence upstream

of hutH from S. griseu s ...... 114

26. Evidence indicating the nature of the mutation in SKK896 127

27. Circular map of pGEM~4Z ...... 141

26. Circular maps of pUC 18 and piJ2925 ...... 143

29. Circular map of plJ702 ...... 145

30. Circular map of plJ486 and plJ487 ...... 147

31. Circular map of pXE4 ...... 149

xi CHAPTER I

Introduction

Histidine Catabolism

The amino acid L-histidine is an essential metabolite in all organisms. L-histidine is incorporated into proteins and may be used as a source of carbon, nitrogen, and energy. Catabolism of L-histidine as a source of carbon and nitrogen in bacteria is most commonly initiated by histidine ammonia-lyase (EC 4.3.1.3; histidase), the product of the hutH gene. As the first enzyme in the histidine utilization (hut) pathway, histidase catalyzes the non-oxidative deamination of L-histidine to urocanate and ammonia (Fig.

1). In bacteria such as Bacillus subtilis (Chasin and Magasanik, 1968) and the enteric bacteria Klebsiella aerogenes (Lund and Magasanik, 1978) and Salmonella typhimurium

(Magasanik, 1978), 1 mol of L-histidine is degraded via a four-enzyme pathway to 1 mol each of ammonia, formamide, and glutamate. In contrast, in other bacteria such as the pseudomonads (Coote and Hassall, 1973; Lessie and Neidhardt, 1967) and streptomycetes (Kendrick and Wheeiis, 1982; Kroening and Kendrick, 1989), 1 mol of L- histidine is catabolized by five enzymatic reactions to 2 mols of ammonia and 1 mol of both glutamate and formate. In either pathway, the first three steps are identical and are catalyzed by histidase, urocanase, and imidazolonepropionate hydrolase. Ammonia can then serve as the nitrogen source, and glutamate can enter into central metabolism.

1 Fig. 1. Pathways of L-histidine utilization in bacteria. Pathway 1 occurs in Bacillus

and the enteric bacteria, pathway 2 in the pseudomonads and

streptomycetes.

2 H H C-C-COOH H KlH,

L-Histidine

H -C=C-CO OH i H *vi H ,0 Urocanic Acid urocanase H H

Imidazolonepropionic Acid

H H H tmldazoloneproplonate HOOC — C -C -C -C O O H hydrolase I H H NH H ,0 formlmlnoglutamate HN = C H Imlnohydrolase 2 N-Formimino-L-Glutamic Acid H H H formlmlnoglutamcrte H20 HOOC — C -6 -C -C O O H fbrmlmlnonydrolase Y HN H lH

H A O HCONH N-formylglutamic Acid

formylglutamase H H H HOOC — C-b-C-COOH N H H iHCOOH H2 L-Glutamic Acid

Figure 1 Formamide is thought to be excreted (Kaminskas et al., 1970; Lund and Magasanik,

1965; Neidhardt and Magasanik, 1957). The pathway of L-histidine catabolism in

mammals is identical to that in B. subtilis except in the last step, in which

formiminoglutamate is hydrolyzed by glutamate formiminotransferase to

formiminotetrahydrofolate (Levy, 1989). All of the enzymes for histidine catabolism are

found in the liver and epidermis. Formiminotetrahydrofolate can then be converted to

formyltetrahydrofolate, a biologically active form of folate coenzyme (Levy, 1989).

Biochemical Analysis of Histidase

L-histidine contains amino, carboxyl, and imidazole groups. Inhibition studies

have suggested that histidase interacts with these three groups of L-histidine at the active

site (Givot et al., 1969). D-histidine, which contains functional groups corresponding to

the three groups of L-histidine, can serve as a potent competitive inhibitor, while

histamine, containing only two functional groups, is a poor Inhibitor (Givot et al., 1969).

Furuta et al. (1992) proposed an elimination reaction in which the a-amino group of

L-histidine associates with histidase and forms a carbanlon intermediate. Urocanate first

dissociates from the carbanion intermediate, and subsequently the amino group is

released from histidase to form ammonia.

Histidase from 8. subtilis (Magasanik et al., 1971) and Pseudomonas species

(Consevage and Phillips, 1985; K.E. Kendrick, unpublished), displays linear reaction

kinetics. Interestingly, streptomycete histidase shows hysteresis (Kendrick, 1979;

Kroening and Kendrick, 1987; Roos et al., 1992). Kroening and Kendrick (1987) observed that activation of histidase from Streptomyces griseus occurred during the 5

course of the enzymatic reaction. The enzyme was twice as active after 10 minutes of

reaction when crude extract prepared from vegetative mycetia of S. griseus was the

source of enzyme. Both maximal histidase activity and linear reaction Kinetics were

achieved by treatment of this crude extract at 60°C for 10 min. Whenever L-histidine was

present in the culture, the total levels of histidase from exponentially growing mycelia or

from a culture in the stationary phase were virtually identical, but a much lower level of

active enzyme was detected in unheated extracts prepared from the cultures that had

entered stationary phase. It appears that the activity of histidase from S. griseus is

modulated by a mechanism that leads to growth phase-dependent fluctuations in

histidase activity (Kroening and Kendrick, 1987}.

There are two forms of histidase from 5. griseus, an active form and an inactive

form (P.J. White, unpublished). There are no differences in molecular weight, subunit

organization (tetrameric), and isoelectric point between the two forms of purified

histidase. The inactive form of histidase can be converted to the active enzyme in vitro

by a partially purified protein (HAP, histidase-activating protein) from S. griseus or by

alkaline phosphatase. Because the activating fraction did not hydrolyze p-

nitrophenylphosphate and did not co-purify with a protein that did hydrolyze that

substrate, the activating protein is probably not streptomycete alkaline phosphatase. This

result suggests that histidase activation may be governed by a factor that catalyzes the

hydrolysis of a phosphoryl group.

Inactivation of pseudomonad histidase by K14CN or NaB9H4 and subsequent identification of [14C]aspartate or [3H]alanine, respectively, following acid hydrolysis of 6

labeled histidase indicated that dehydroalanine is required for catalysis (Consevage and

Phillips, 1985; Wickner, 1969). An analogous enzyme, phenylalanine ammonia-lyase (EC

4.3.1.5), which has been purified from plants (Havir and Hanson, 1975) and yeast

(Hodgins, 1971), is also thought to contain dehydroalanine at the active site. Similar

experiments using histidase purified from Pseudomonas sp. ATCC 11299b with labeled

iodo[14C2]acetate as a probe identified a cysteinyl residue at the active site (Klee and

Gladner, 1972). Histidase from S. griseus is inactivated by cyanide (Wu et al., 1992),

which is thought to act by interacting with dehydroalanine (Consevage and Phillips,

1985). White and Kendrick (1993) also showed that S. griseus histidase is inactivated by

dicarbonyl reagents, which react specifically with arginyi residues in proteins (Patthy and

Thesz, 1980). Histidase from P. putida, which had been labeled with [14C] serine during

biosynthesis, was treated with KCN followed by acid hydrolysis. The hydrolyzed histidase

did not contain the predicted labeled aspartate, suggesting that serine is not the

precursor for dehydroalanine (Consevage and Phillips, 1985). In contrast, a recent report

by Hernandez et al. (1993) demonstrated that a modified serine residue (Ser143 of P. putida histidase) is likely to be involved in catalysis. Although the chemical structure of the modified serine was not determined, the molecular mass of the modified serine (184

Da) does not correspond to that of dehydroalanine (88 Da). Analysis of amino acid sequences of four histidases as well as three phenylalanine ammonia-lyaaes showed that a region containing the modified serine is conserved among all seven enzymes

(Hernandez et al., 1993). The identification of the chemical nature of serine-143 in the active enzyme will resolve whether dehydroalanine, serine, or another modified form of serine is present at the active site. To date, the amino acid residues at the active site of histidase have not been identified with certainty, and both the origin and the nature of the 7 linkage of dehydroalanine are unknown. Comparisons of histidase primary structures from a variety of organisms should contribute to the identification of the amino acid residues at the active site.

Histidinol phosphate has been identified as a competitive inhibitor of 5. griseus histidase and protects the enzyme from inactivation by cyanide (Wu et al., 1992) and dicarbonyl reagents (White and Kendrick, 1993). There is no report that histidinol phosphate inhibits other histidases. Because both phosphorylated and carboxylated substrates are known to react with arginyl residues in proteins (Patthy and Thesz, 1960), an arginyl residue in S. griseus histidase probably interacts with both histidinol phosphate and L-histidine. This residue is likely to be essential for catalysis.

The identification of amino acid residues required for histidase catalysis is also of clinical interest. Histidinemia, a hereditary disorder in mammals, results from the accumulation of histidine in body fluids due to histidase deficiency (Levy, 1989).

Genetic Analysis of Histidine Utilization

In all bacterial species examined to date, the genes of histidine utilization are clustered on each genetic map. Hut mutant analysis has shown that the hut genes are organized in a single operon in B. subtilis (Fig. 2; Chasin and Magasanik, 1968; Kimhi and Magasanik, 1970; Oda et al., 1988), whereas in Pseudomonas putida (Consevage et al., 1985; Hu and Phillips, 1988), the hut genes are divided into four adjacent transcriptional units (Rg. 2). Analysis of the nucleotide sequence of the hutH gene from

P. putida has revealed an additional potentially weak promoter immediately upstream of Fig. 2. Organization and transcriptional units of the bacterial hut genes. The

arrows represent the transcriptional units and the directions of the

transcripts. Question marks indicate potential transcripts that have not

been verified. A region containing a stem-and-loop structure between

hutP and hutU in B. subtilis is indicated. The gene products are HutC, a

repressor protein; HutF, formylglutamate hydrolase; HutG,

formiminoglutamate iminohydrolase in P. putida and formiminoglutamate

formiminohydrolase in other bacteria; HutH, histidase; Hutl,

imidazolonepropionate hydrolase; HutP, a positive regulator; and HutU,

urocanase. The hut genes are not drawn to scale.

8 9

H U

B. subtilis

U

? - ► K. aerogenes ►

S. typhimurium

H U

< ■ P. putida

Figure 2 10 the hutH structural gene, but whether the hutH gene is transcribed from this putative promoter is stilt unknown (Consevage and Phillips, 1990). The hut genes are arranged in two and three operons on the Klebsiella aerogenes and Salmonella typhimurium chromosomes, respectively, between gal and bio (Goldberg and Magasanik, 1975;

Magasanik, 1978). Whether the hutC gene, encoding the hut repressor, is also part of the hudGC operon in K. aerogenes remains uncertain (Schwacha and Bender, 1990). In all of these cases, hutH is in an operon and is co-transcribed with hutU.

In the enteric bacteria, pseudomonads, and Bacillus, synthesis of the hut enzymes is inducible (Brill and Magasanik, 1969; Chasin and Magasanik, 1968; Coote and Hassall,

1973; Hu et al., 1987; Schlesinger et al., 1965) and generally subject to catabolite repression (Brill and Magasanik, 1969; Chasin and Magasanik, 1968; Coote and Hassall,

1973; Leidigh and Wheeiis, 1973; Prival and Magasanik, 1971) and nitrogen regulation

(Magasanik, 1978; Prival and Magasanik, 1971) a! the transcriptional level. In B. subtilis, expression of the hut genes is positively controlled by HutP, a diffusible positive regulator

(Oda et al., 1988). A mutant of B. subtilis defective in hutP lacks all four hut enzymes, and the mutant containing hutP in trans shows full production of the hut enzymes. A region that contains a stem-and-loop structure (Fig. 2) lying between hutP and hutH has been proposed to function as a transcription terminator. Oda et al. (1988) proposed that in the presence of the inducer, L-histidine, the HutP protein may associate with this stem- and-loop structure so that transcription initiated at the hut promoter can proceed beyond this stem-and-loop region into the neighboring hut structural genes. 11

Negative regulation mediated by HutC, a repressor protein, occurs in the enteric

bacteria (Magasanik, 1978) and P. putida (Hu et al., 1989). Urocanate is the

physiological inducer of the hut enzymes in the enteric bacteria (Brill and Magasanik,

1969; Schlesinger et al., 1965) and pseudomonads (Hu et al., 1989). In P. putida, hutG

expression is also induced by formylglutamate, the substrate of HutG (Hu et al., 1989).

The hutC genes from K. aerogenes (Schwacha and Bender, 1990) and P. putida (Allison

and Phillips, 1990) have been cloned and sequenced. There is 62% amino acid

sequence identity between the repressors from these two organisms. Regions near the

amino terminus of HutC from these two bacterial species show properties of the helix- tum-helix motif found in many DNA-binding proteins such as the lac repressor protein

and the catabolite activator protein from E. coii (Schwacha and Bender, 1990).

Examination of the partially purified P. putida repressor protein for its ability to bind various restriction fragments spanning the entire coding region of the hut genes has localized three repressor binding sites (Hu et al., 1989). These three sites are located in the vicinity of regions preceding hutF (and hutC\ see below), hutU, and hutG, providing evidence that the hut operons in P. putida contain four transcriptional units. Interestingly, only one repressor binding site between hutC and hutF was identified. The researchers proposed that a single regulatory region is shared by two genes which are transcribed in opposite directions (Fig. 2; Hu et al., 1989). Determination of the nucleotide sequence between the hutC and hutF coding regions may resolve this question. There is considerable reduction of repressor binding ability at all three sites in the presence of urocanate but not in the presence of L-histidine (Hu et al., 1969). Either formylglutamate, the substrate of HutG, or urocanate can inhibit binding of the same repressor protein to 12

the site preceding hutG (Hu et al., 1989). These results support the physiological data

that in P. putida, urocanate and formylglutamate, rather than L-histidine, are the effective

inducers for the hut operons and hutG, respectively.

By gel retardation and footprinting analyses, a region containing approximately

40 bp spanning positions -10 to -50 relative to the transcription initiation site for hutU in

P. putida was identified as the repressor binding site (Hu et al., 1989). This region

contains an inverted repeat of 16 nucleotides spanning the -35 position. Similar

nucleotide sequences in inverted repeat orientation were also found in the regions

preceding hutC in P. putida (Allison and Phillips, 1990) and K. aerogenes (Schwacha and

Bender, 1990) as well as the hutUH operon in K. aerogenes (Schwacha and Bender,

1990). These inverted repeats are likely to serve as repressor recognition sequences.

Thus, in P. putida and K. aerogenes, the mechanism of repression is expected to prevent

access of RNA polymerase to the promoters of the hut genes by the repressor, and hutC

appears to be autoregulated.

Catabolite repression of the hut genes in K. aerogenes (Nieuwkoop et al., 1984;

Osuna et al., 1991) and S. typhimurium (Brill and Magasanik, 1969) is mediated by

catabolite activator protein (CAP) and cyclic AMP (cAMP). In S. subtilis and

pseudomonads, the molecular basis of catabolite repression is unknown but appears to

be independent of cAMP. In Pseudomonas aeruginosa, the level of intracellular cAMP

remains relatively constant during growth on various carbon sources, and repression of the hut genes cannot be relieved by the addition of exogenous cAMP (Phillips and

Mulfinger, 1981). AS. subtilis mutant that is insensitive to catabolite repression has been 13 mapped to a locus near hutP (Chasin and Magasanik, 1968). DNA fragment deletion analysis revealed a region containing an inverted repeat of 27 nucleotides within the huff* coding sequence that Is required for catabolite repression (Oda et al., 1992). The nucleotide sequence of this inverted repeat resembles those required for catabolite repression of amyE and the gnt operon in the same organism (Oda et al., 1992).

In K. aerogenes, a DNA sequence centered at position -82 relative to the transcription initiation site of the hutUH operon is responsible for catabolite repression

(Osuna et al., 1991). In vitro transcription analysis showed that two transcripts of unknown function transcribed in opposite direction to that of the hutUH operon were predominant over the hutUH mRNA (from promoter hutUp) during conditions that exerted catabolite repression (Osuna et al., 1990). These two transcripts, originating from two overlapping promoters, Pc, and PCj, initiated at -70 and -80 relative to the transcription initiation nucleotide of the hutUH operon. In the presence of exogenous CAP, cAMP, and

RNA polymerase purified from K. aerogenes, synthesis of these two transcripts is repressed and hutUp is active. Thus, expression of the hutUH operon in K. aerogenes is proposed to be activated by a mechanism in which: (1) occupancy of the CAP-cAMP complex at Pc, and PCj hinders the preferential binding of RNA polymerase at these two promoters, thereby favoring the binding of RNA polymerase at hutUp\ (2) the CAP-cAMP complex serves as a positive activator for enhancement of RNA polymerase binding at hutUp, or (3) the CAP-cAMP complex plays both direct and indirect roles as described above in the activation of hutUp (Osuna et al., 1991). 14

Synthesis of the hut enzymes in K. aerogenes (Prival and Magasanik, 1971) and

pseudomonads (Potts and Ciarke, 1976) but not in B. subtilis (Chasin and Magasanik,

1968) and S. typhimurium (Goldberg et al., 1976) is activated upon nitrogen limitation.

Activation of histidase synthesis in response to nitrogen limitation in K. aerogenes

requires the nitrogen regulatory system (Ntr) and nitrogen assimilation control (nac) gene.

Mutants of K. aerogenes with defects in nac, the Ntr system, or Ntr-dependent RNA

polymerase a94 failed to produce histidase when grown in a nitrogen-limiting medium

(Macaluso et al., 1990). Analysis of the nucleotide sequence of nac revealed consensus sequences specifying a o^-dependent promoter and an NtrC (also called NR|f a transcriptional activator) binding site (Schwacha and Bender, 1993a). The deduced amino acid sequence of Nac, the product of nac, is similar to those of the LysR family of regulatory proteins, suggesting that Nac is a DNA-binding protein and its synthesis is regulated by the Ntr system. Furthermore, when IPTG was present, cells carrying the nac gene fused to the IPTG-inducible lac promoter rather than the original Ntr-activated promoter produced histidase (Schwacha and Bender, 1993b). Thus, Nac alone without an active Ntr system is sufficient to activate histidase synthesis.

A model explaining hut gene expression by nitrogen regulation in K. aerogenes has been proposed by Bender and coworkers (Schwacha and Bender, 1993b).

According to this model, the Ntr system activates nac in response to nitrogen limitation, and Nac activates hut gene expression. Whether Nac directly activates the hut operons or acts through another system is still unknown. When nac was placed under the control of the IPTG-inducible lac promoter, Nac not only activated many other nitrogen-regulated systems such as the proline utilization system {put) and urea degradation system, both 15

of which provide ammonia, but also repressed the synthesis of glutamate

dehydrogenase, which is required for ammonia assimilation. Interestingly, activation of

histidase synthesis requires a higher level of Nac than does activation of urease

synthesis. Thus, when grown in a medium containing both urea and L-histidine, cells will

utilize urea as nitrogen source and target L-histidine for biosynthesis (Schwacha and

Bender, 1993b).

In contrast to K. aerogenes, the hut genes in 5. typhimurium are not subject to

nitrogen regulation. When present in the cytoplasm of K. aerogenes, however, the hut

genes of S. typhimurium are activated in response to nitrogen starvation (Goldberg et al.,

1976), suggesting that a factor required for activation of the hut genes is absent from S.

typhimurium. An experiment to determine the function of nac from K. aerogenes in S. typhimurium may explain this observation.

Although derepression of Hut enzyme synthesis was not observed during nitrogen-limiting growth in B. subtilis (Chasin and Magasanik, 1968), synthesis of several other B. subtilis nitrogen-degradative enzymes such as urease and asparaginase is activated by nitrogen limitation (Atkinson et al., 1990). Fisher and coworkers (Atkinson et al., 1990) have shown that expression of the hut genes in B. subtilis is repressed in the presence of a mixture of 16 amino acids (lacking L-histidine), and this repression is independent of carbon catabolite repression and nitrogen regulation of glutamine synthetase. Thus, it is apparent that the amino acid repression of the hut operon in B. subtilis is mediated by a mechanism that is distinct from those that regulate catabolite 16 repression, the synthesis of glutamine synthetase, and production of several other nitrogen-degradative enzymes.

Biological Importance of Streptomycee

Streptomycetes are gram-positive soil bacteria that, upon nutrient deprivation, undergo a developmental process in which vegetatively growing mycelia begin to form cross walls and become spores (Fig. 3; Chater, 1964). This process produces hundreds of spores from a single mycelial unit. Streptomycetes produce the majority of commercially important secondary metabolites such as antibiotics (Chater, 1964).

Because synthesis of secondary metabolites and sporulation often occur simultaneously

(Hopwood, 1986), streptomycetes are useful as a model system to study the relationship between cell differentiation and secondary metabolism. Kendrick and Ensign (1983) have developed a system in which vegetative cells of 5. griseus can be induced to sporulate rapidly and synchronously in submerged culture. This induction system involves transferring mycelia from a rich medium to a defined medium or to a phosphate- or nitrogen-limiting medium. This system permits the study of molecular and biochemical changes during sporulation on a useful scale, which has not yet been developed for the genetically best characterized species of streptomycete, 5. coelicolor.

Nitrogen Metabolism in Streptomycee

Very little is known about the metabolism of nitrogenous compounds in

Streptomyces. Most of the work has focused on enzymes involved in secondary metabolism. Although nitrogen regulation occurs in streptomycetes, histidase is not subject to this regulatory mechanism (Ba$car&n et al., 1989; Kendrick and Wheelis, 1982; Fig. 3. Life cycle of Streptomyces. A, free spore#; B, germinating spores; C,

vegetative mycelia; D, reproductive spores.

17 18 r

D B

Figure 3 19

Kroening and Kendrick, 1987). In contrast, studies of nitrogen-deregulated mutants of

S. clavuligerus established that glutamine synthetase (GS), urease, arginase, and

ornithine aminotransferase are regulated in response to the availability of ammonia

(Bascar&n et al., 1989). GS has been the focus of several studies. Assimilation of

ammonia by the glutamine synthetase-glutamate synthase (GOGAT) pathway, when the

concentration of ammonia is low, or the glutamate dehydrogenase (GDH) pathway, when

ammonia is abundant, has been demonstrated in Streptomyces. GS activity was

detected in S. coelicolor (Fisher and Wray, 1989), Streptomyces cattieya (Paress and

Streicher, 1985), Streptomyces clavuligerus (Braha et al., 1986), and Streptomyces

venezuelae (Shapiro and Vining, 1983). Both S. coelicolor and S. venezuelae contain

GOGAT and GDH. However, S. clavuligerus appears to produce GOGAT but not GDH.

GS is regulated transcriptionally and post-translationaily. In S. coelicolor (Fisher

and Wray, 1989), transcription of the GS structural gene, glnA, is regulated in response to the availability of ammonia. The glnA gene is positively regulated by glnR in S. coelicolor (Wray et at., 1991). The deduced amino acid sequence of glnR is similar to those of bacterial response regulators such as OmpR and PhoP, suggesting that the

GlnR protein is a transcriptional activator (Wray and Fisher, 1993). Additionally, the active form of GS becomes rapidly inactivated by adenylylation in response to an increase in the concentration of ammonia in S. cattieya (Streicher and Tyler, 1981) and S. coelicolor

(Fisher and Wray, 1989). in contrast, there is no significant change in the level of GS activity in S. venezuelae when grown in various nitrogen sources (Shapiro and Vining,

1983). 20

Both S. hygroscopicus (Kumada et al., 1990) and S. viridochromogenes

(Behrmann et al., 1990) contain a second GS structural gene that encodes a distinct GS isoform (GSII). The physiological role of GSII is still unclear. The amino acid sequence and thermostability of GSII are closely related to those of eucaryotic GS. Southern analysis suggested that multiple GS genes may be common in Streptomyces (Behrmann et al., 1990; Kumada et al., 1990)

Streptomycetes produce a variety of extracellular hydrolytic enzymes such as proteases (Chandrasekaran and Dhar, 1987; Narahashi, 1970) that degrade extracellular macromolecules to oligomers. Presumably these enzymes serve to generate nutrients in the soil environment. The oligomers are transported and metabolized to support growth, cell differentiation, and biosynthesis of secondary metabolites. The amino acid

L-histidine is likely to derive from the extracellular proteins in the Streptomyces natural environment

When L-histidine is the sole source of nitrogen, it must supply all of the nitrogen required for growth, morphogenesis, and secondary metabolism. For example, as sole nitrogen source, L-histidine must be converted into protein, the nitrogenous antibiotics cycloheximide and streptomycin (produced by the wild-type strain of S. griseus), and all other nitrogenous compounds necessary for growth and survival (Fig. 4). In these respects, S. griseus must be capable of controlling the flux of L-histidine to ensure the adequate supply of precursor for each process. One way in which the flux of L-histidine could be controlled would be by tightly regulating the activity of a key enzyme in L- Fig. 4. Metabolism of L-histidine in streptomycetes.

21 22

protein

amino acids transport ■=> L-histidine NH biosynthesis antibiotics urocanate other nitrogenous compounds

glutamate

Figure 4 23 histidine metabolism. Such an enzyme would be expected to be a branch-point enzyme whose activity may be regulated in a hysteretic fashion (Frieden, 1979). Kroening and

Kendrick (1987) have shown that in S. griseus a rapid decrease in histidase activity occurs at the onset of sporulation, and histidase activity is restored upon transferring the sporulating culture to a nutritionally replete medium. Bormann et al. (1989) showed that

L-histidine is a direct precursor of the nucleoside peptide antibiotic nikkomycin, which is produced by S. tendae. Histidine aminotransferase, which converts L-histidine to imidazole pyruvate (a precursor of nikkomycin), can be detected only in a Hut' mutant of S. tendae that lacks histidase activity, and production of nikkomycin in the mutant is unimpaired. These two examples suggest that histidase may be a branch-point enzyme in streptomycetes.

Enzymological studies of streptomycete histidase have shown that this enzyme has characteristics distinct from other bacterial and mammalian histidases. Hysteretic behavior indicative of an activation event was observed in streptomycete histidase in crude extracts (Kendrick, 1979; Kroening and Kendrick, 1987; Roos et al., 1992) as well as in partially purified protein preparations (P.J. White, unpublished). Unlike pseudomonad histidase (Consevage and Phillips, 1985; Rechler, 1969), which is activated by thiol reagents and divalent cations, streptomycete histidase Is unaffected by these chemicals (Wu et al., 1992). The of purified histidase from 5. griseus for L-histidine is 0.6 mM (Wu et al., 1992), which is approximately 5- to 20-fold lower than the affinity constants of other bacterial histidases (Consevage and Phillips, 1985; Dalen, 1973; Klee et al., 1975). Histidinol phosphate is a strong competitive inhibitor of streptomycete 24

histidase; at best, it only weakly inhibits pseudomonad histidase (K.V. Srinivasan,

unpublished).

In the enteric bacteria (Magasanik, 1978), pseudomonads (Hu et al., 1987), and

Bacillus subtilis (Chasin and Magasanik, 1968; Oda et al., 1992), synthesis of histidase

is subject to induction, carbon catabolite repression, and nitrogen regulation at the level

of transcription. The synthesis of streptomycete histidase is controlled by neither carbon

catabolite repression nor nitrogen regulation (Kendrick and Wheelis, 1982; Kroening and

Kendrick, 1987): histidase activity can be detected whenever L-histidine is present in the

growth medium regardless of the presence of other carbon and nitrogen sources. It

appears therefore that the regulation of histidase synthesis in streptomycetes is also distinct from that in other bacteria.

Goals of the Project

To determine the regulation of hutH expression in S. griseus, I first set out to clone the hutH gene from S. griseus by plaque hybridization with an oligonucleotide probe designed from the amino acid sequence of the N-terminus of purified histidase from S. griseus. The hutH gene served as the foundation for the following goals to extend our understanding of the hut system in this organism.

1. Determination of the hutH nucleotide sequence.

2. Determination of the organization of the hutH gene.

3. Determination of the mechanism of regulation of hutH expression. 25

4. Identification of the defect in a Hut mutant strain that is deficient in

histidase activity.

The results of this study show that there are unique features of the structure, organization, and regulation of hutH in S. griseus. The deduced amino acid sequence of histidase from S. griseus is similar to that of other bacterial and mammalian histidases but is phylogenetically more distant. Unlike hutH expression in other bacteria, the hutH transcript in 5. griseus is monocistronic. Transcription and translation of hutH in S. griseus initiate at the same nucleotide position. Evidence suggests that hutH expression is controlled by a negative regulatory factor that operates a considerable distance upstream of the promoter. These unique characteristics indicate that expression of the hutH gene in S. griseus is subject to distinct regulatory mechanisms. CHAPTER II

MATERIALS AND METHODS

I. Bacterial strains, plasmids, and bacteriophage

The bacterial strains used in this study are listed in Table 1. The wild-type strain of Streptomyces griseus, SKK821, was obtained from the Northern Regional Research

Laboratory, Peoria, IL, as NRRL B-2682. All of the S. griseus strains listed below are derivatives of SKK821. Strain Rfs2 was isolated by R. L. Sharp in this laboratory by curing a transformant of SKK821 that contained plJ702 from 5. IMdans strain TK24. Rfs2 exhibits a ten-fold increase in transformation frequency compared with that of the parent strain. The phenotypes of strain SKKB32, a histidine auxotroph, and SKK896, a histidine utilization mutant (Hut), were described previously (Kroening and Kendrick, 1989). The

E. coli strains and S. IMdans strain TK24 were used for the construction, manipulation, and propagation of plasmids. S. coelicolor M145 was used in Southern hybridization studies.

Table 2 lists all of the plasmid vectors and phage used in this study. The structures of the plasmid vectors are presented in Appendix A. The methods of construction of plasmids derived from these vectors are described in Results, and detailed descriptions of the recombinant plasmids are provided in Appendix B.

26 27

Table 1. Bacterial strains used.

Strain Relevant Genotype Source or Reference

S. griteu* strains NRRL B-2682 (SKK821) wild-type NRRL* Rfs2 restriction-defective this lab I SKK832 hisG1 (Kroening and Kendrick, 1989) SKK896 hut-2 (Kroening and Kendrick, 1969) Other Streptomyces strsins S. coelicolor M145 wild-type M.J. Bibb | S. Irvidans TK24 wild-type D.A. Hopwood E co il strains

LE392 — Promega

TB1 — C.J. Daniels

NRRL, Northern Regional Research Laboratory, Peoria, IL 28

Table 2. Plasmid vectors and bacteriophage used.

| Host System Plasmid or Relevant Source Phage Phenotype*

E. coli/Streptomyces pXE4 XylE (F), Tsr, Amp J. Westpheiing (bifunctional) £. coli PGEM-4Z* Amp Promega plJ2925° Amp G.R. Janssen pUC18 Amp C.J. Daniels

\ E. coli AEMBL4 clM7 Promega | Streptomyces plJ486 Km (F), Tsr D.A. Hopwood plJ487 Km (F), Tsr D.A. Hopwood plJ702 Mel, Tsr D.A. Hopwood

B Amp, ampicillin resistance; c l^, temperature-sensitive repressor; Km, kanamycin resistance (aminoglycoside 3 -phosphotransferase II); Mel, tyrosinase; F, promoterless;

Tsr, thiostrepton resistance; XylE, catechol 2,3-dioxygenase. b a transcription vector containing SP6 and T7 promoters. c a derivative of pUCid containing additional restriction sites in the polylinker region. 29

II. Media and culture conditions

The media used in this study and their purposes are described in Table 3. With

the exception of minimal agar medium, the compositions and/or uses of the individual

media are found in the indicated references. Minimal agar medium contained (% wt/vol):

0.05% KaHPO*, 0.02% MgS04-7Ha0, 0.001% FeS04*7Ha0, 20mM L-glutamate, pH 7.0-

7.2, and 1 % Noble agar (Difco Laboratories, Detroit, Ml). Trypticase soy broth (TSB) was

purchased from BBL, Microbiology Systems, Cockeysville, MD.

When required for screening E. coli transformants, the color indicator 5-bromo-4-

chloro-3-indolyl-/f-D-galactoside (X-gal; BRL, Bethesda Research Laboratories Inc.,

Gaithersburg, MD) at a concentration of 40 pg/ml was included in LB agar plates. When

S. griseus SKK832 and its derivatives were grown, 0.1 mM L-histidine (USB, United States

Biochemicals Corp., Cleveland, OH) was provided to satisfy the requirement of the

auxotroph.

Kanamycin sulfate (Sigma Chemical Co., St. Louis, MO) was used in the

promoter-probe studies. Other antibiotics were used at the following concentrations for

culture maintenance and selection. Ampicillin (sodium salt; Sigma) was included at 50

lig/ml for pXE4 derivatives and 100 ng/ml for pUC18 derivatives. Thiostrepton (Squibb

Institute for Medical Research, Princeton, NJ, or Sigma) was included in liquid media at

5 |ig/ml. Thiostrepton was provided at 5 jig/ml for S. griseus cultures on solid media, at

10 pg/ml for S. IMdans containing pXE4 derivatives, and at 40 pg/ml for S. IMdans containing high copy number plasmids. 30

Table 3. Media used.

Medium Purpose Strain Reference | 2XYT genomic DNA isolation S. griseus (Sambrook et I al., 1989) CAA medium analysis of Hut enzymes, S. griseus (Kendrick and promoter probe studies Ensign, 1983) using pXE4, and RNA isolation 1 Glutamate analysis of Hut enzymes, S. griseus (Kendrick and 1 minimal promoter probe studies Ensign, 1983) I medium using pXE4, and RNA isolation LB (Luria broth) plasmid isolation and E. coli (Sambrook et propagation of phage al., 1989) LB agar strain maintenance E. coii (Sambrook et al., 1989) Minimal agar promoter probe studies S. griseus and described in medium using plJ486 and plJ487 S. IMdans text R2YE spore preparations, S. coelicolor (Hopwood et | protoplast regeneration, and and S. IMdans al., 1985) | strain maintenance SpM liquid spore preparations S. griseus (Kendrick and | Wheelis, 1982) SpM agar strain maintenance S. griseus (Kendrick and 1 Wheelis, 1982) I SpmR regeneration of protoplasts S. griseus (Babcock and D Kendrick, U 1988) Terrific broth phage propagation E. coli LE392 (Babcock and Kendrick, 1988) TSB protoplast generation and S. coelicolor, (Babcock and | (Trypticase Soy isolation of plasmid DNA $. griseus, and Kendrick, | Broth) S. IMdans 1988) | 31

Unless otherwise stated, cultures of Streptomyces and E. coli were grown at 30°C

and 37°C, respectively. Liquid cultures of Streptomyces strains were grown in flasks in

a rotary shaker at 250 rpm and contained 5% (S. griseus) or 15% (S. coelicolor and S.

IMdans) polyethylene glycol 8000 (PEG-8000; molecular weight, 8000; Sigma) and a coiled spring to improve the dispersion of the mycelia (Kendrick and Wheelis, 1982). In

liquid cultures in a test tube, the PEG and the spring were replaced by a 6.0 mm

diameter glass bead to enhance dispersed growth (P. Bartel, personal communication).

Either liquid SpM or sporulation broth was used to grow cultures of S. griseus for spore preparations, which were prepared as described by Kendrick and Ensign (1983).

Spore suspensions of S. coelicolor and S. IMdans were prepared by harvesting spores from confluently sporulated cultures on R2YE agar after 5 days of incubation. After the addition of 5 ml of sterile distilled H20 to the plate, the spores were scraped off with a pipette, decanted, centrifuged, and washed once with an equal volume of sterile distilled

H20. The spore suspensions were stored in sterile distilled HaO at 4°C. Heat-activated spores pregerminated in TSB (Kendrick and Ensign, 1983) were used as inoculum for liquid media in flasks. A single sporulated colony served as inoculum for liquid cultures in test tubes.

III. Recombinant DNA techniques

A. Isolation of DNA

For propagation of phage AEMBL4, E. coli LE392 was grown in Terrific Broth. A phage lysate was combined with 1 ml of overnight culture of E. coli (1 X 10'° cells/ml) at a multiplicity of infection of 0.01. After 30 min of incubation at 37°C, the infected culture was added to 250 ml prewarmed LB containing 10 mM MgS04-7H20, and incubation was continued for 4*6 hr at 37°C with vigorous shaking until lysis was complete.

Chloroform (5 ml) was added to the lysed culture. Following incubation at 37°C for 30 min, 14.5 g NaCI was added to the culture and the debris was removed by centrifugation at 11,000 x g for 10 min at 4°C. The supernatant was transferred to a flask containing

25 g PEG-8000. The PEG was dissolved by stirring and the suspension was kept on ice overnight at 4°C. The phage particles were collected by centrifugation at 11,000 x g for

10 min at 4°C and suspended in 5 ml SM buffer (0.58% NaCI, 0.2% MgS04-7H20, 0.05

M Tris-HCI, pH 7.5, and 0.01% gelatin [Sambrook et al., 1989]). Purification of phage particles by glycerol gradient centrifugation and extraction of phage DNA were performed as described by Sambrook et al. (1969). The phage DNA was stored in TE (10 mM Tris-

HCI, 1 mM EDTA (disodium salt), pH 8.0) at -20°C.

Cultures of E. coli used for plasmid DNA isolation were grown in 3 ml of LB containing ampicillin. Streptomyces strains were grown in 3 ml of TSB containing thiostrepton, and a glass bead was included in the tube. Plasmid minipreparations were performed according to the method described by Babcock and Kendrick (1988). Plasmid

DNA preparations from both genera were used for restriction mapping, transformation, and DNA fragment isolation. The plasmid minipreparations from E. coli were also used for nucleotide sequence determination. Plasmid DNA to be used for generating nested deletions by exonuclease III or as a template for in vitro transcription was prepared from a 250 ml culture in a 1 L flask, isolated using a scaled-up minipreparation procedure, and purified twice through a cesium chloride density gradient (Hopwood et al., 1985). 33

Streptomyces strains used for genomic DNA isolation were grown in 50 ml of either TSB containing 15% PEG-8000 (S. coelicolor) or 2XYT containing 5% PEG-8000

(S. griseus). The cultures were harvested by centrifugation during early stationary phase for maximal yields. Mycelia were washed once with 10 ml of 3 M sucrose and suspended in 5 ml TE. The cells were lysed by the addition of lysozyme (Sigma) at a final concentration of 0.5 mg/ml and incubated for 5 to 10 min at room temperature. The isolation of genomic DNA from the lysed cells was performed according to the method of Hopwood et al. (1985) except that DNA precipitation by PEG was replaced with precipitation by using 0.1 volume of 3.0 M sodium acetate, pH 6.0, and 2.5 volumes of

95% ethanol. The genomic DNA was stored in TE at -20°C.

B. Generation and manipulation of DNA fragments

Restriction endonucleases were purchased from BRL, Boehringer Mannheim

Biochemicals (Indianapolis, IN), or New England Biolabs (Beverly, MA) and used according to the manufacturer’s recommendations. DNA restriction fragments were resolved by electrophoresis on 1 % horizontal agarose gels submerged in GGB buffer (40 mM Trizma base [USB], 20 mM sodium acetate, 2 mM Na,EDTA (disodium salt), adjusted to pH 8.3 with glacial acetic acid; Babcock and Kendrick, 1988). The resolved DNA fragments were detected by staining with ethidium bromide and photography under UV illumination. Restriction fragments prepared from X DNA or plasmid DNA were used as size standards.

Two alternative methods were used to isolate DNA fragments from agarose gels for use in cloning and as probes in hybridization analyses. After digestion with restriction 34 enzymes, the DNA fragments were resolved on standard agarose gels and stained as described above. SeaKem QTG agarose (FMC BioProducts, Rockland, ME) was used to maximize plasmid ligation. The DNA fragments were excised from the gel and purified by either the phenol-freeze-fracture method (Huff, 1991) or the Prep-A-Gene method (Bio-

Rad Laboratories, Richmond, CA). The yields by using either method were approximately

70%. The fragments were stored in distilled H20 or TE at -20°C. In some cases, after fragment isolation the ends of the restriction fragments were made blunt by using Klenow fragment of E. coti DNA polymerase I (BRL) according to the method of Sambrook et al.

(1969).

Deletion subclones in pUC18 were generated by exonuclease III (Henikoff, 1984).

To generate unidirectional deletions of a 3 kb fragment in pUCl8, CsCI-purified plasmid

DNA (7.5 pg) was digested with Xbal to leave a 5’ overhang and Psft to leave a 3* overhang. The DNA was purified by extraction with phenol-chloroform and two successive extractions with chloroform and precipitated with sodium acetate and ethanol overnight at -20°C. The DNA was collected by centrifugation, washed with 70% ethanol, and dissolved in 75 pi exonuclease III buffer (0.66 mM MgCI2 and 66 mM Tris-HCI, pH

8.0). The deletion reaction was started by the addition of 1 pi of exonuclease III (175 units/pl; Boehringer). The reaction mixture was incubated at 37°C, and aliquots of 4 pl were transferred at intervals to 25 pl of SI nuclease digestion buffer (0.25 M NaCI, 1.0 mM ZnS04, 5% glycerol, and 30 mM potassium acetate, pH 4.6) in a microcentrifuge tube at 4°C. Under these conditions, approximately 350 nucleotides were hydrolyzed in 1 min.

After all aliquots had been transferred, 25 pl S1 nuclease buffer (50 mM sodium acetate, pH 5.7, 200 mM NaCI, 1mM ZnS04, and 0.5% glycerol) containing S1 nuclease (BRL) at 35 a final concentration of 0.5 u/pl was added to each tube. The samples were incubated

at room temperature for 30 min, and the reaction was stopped by adding 6 pl S1 stop

buffer (125 mM EOTA (disodium salt) and 0.5 mM Tris-HCI, pH 8.0). The DNA from each sample was extracted and precipitated as described above. The DNA was recovered by centrifugation, washed with 70% ethanol, and dissolved in 40 pl T4 ligase buffer (BRL).

After the addition of 1 pl T4 DNA ligase (1 unit/pl; BRL), the ligation mixtures were left for

2-3 hr at room temperature. The mixtures were placed at 65°C for 10 min to inactivate the ligase and then digested with Pst\ or Xbal to ensure that the original plasmid DNA would be unavailable for transformation. The ligation mixtures were used to transform competent E coli TB1 cells.

To achieve optimal conditions for plasmid ligation, the molar ratio of insert fragment to vector was maintained at 3 to 1. The vector was dephosphorylated to minimize self-ligation and maximize ligation to an insert fragment. After digestion with the restriction enzyme, E coli vectors and Stroptomyces vectors still in the digestion mixtures were treated with alkaline phosphatase (Boehringer) at a concentration of 1 unit per pg of DNA at 37°C for 20 min and 30 min, respectively. The reactions were transferred to

70°C for 10 min to inactivate the alkaline phosphatase. The DNA was extracted and precipitated as described above. The DNA was recovered by centrifugation, washed with

70% ethanol, and dissolved in distilled HaO.

Oligonucleotides were synthesized on an Applied Biosystems model 380B DNA synthesizer (Biochemical Instrument Center, The Ohio State University). Oligonucleotides

(0.1 pmol in 2.5 ml solution) were first deblocked to be suitable for labeling. A screw- 36

capped microcentrifuge tube containing 0.3 ml of the oligonucleotide solution was

incubated at 55°C for 12-17 hr. The solution was evaporated in a centrifuge evaporator

(SpeedVac, Savant Instruments Inc., Farmingdale, NY). When the volume of the solution

was reduced to less than 20 pl (2-3 hr), 0.5 ml distilled H20 was added and the solution

was again evaporated. The water wash was repeated twice. After the final wash, the

oligonucleotide was precipitated with 0.5 volumes of 7.5 M ammonium acetate and 2.5

volumes of 100% ethanol and kept at -20°C for 2 hr. The oligonucleotide was recovered

by centrifugation, washed with ethanol, dried, and dissolved in distilled H20. The

concentration of the oligonucleotide was determined spectrophotometrically and adjusted

to 100 pmol/pl with distilled H20. The oligonucleotide solution was stored at -20°C.

The oligonucleotides were end-labeled with T4 polynucleotide kinase (BRL) and

y-(“ P)-ATP (Amersham Corp., Arlington Heights, IL or ICN Biomedicals, Inc., Irvine, CA)

as described by Sambrook et al. (1969). The oligonucleotides labeled at their 5' ends

with were used as probes in Southern and plaque hybridizations and also as primers

in generating sequencing ladders in high resolution S1 mapping and in primer extension

analysis.

For labeling at their 5’ ends, the appropriate DNA restriction fragments containing

5’ overhanging ends were first dephosphorylated with calf intestine alkaline phosphatase

(Boehringer), then treated with T4 polynucleotide kinase as described by Sambrook et al. (1989). The fragments were used as probes in S1 mapping studies. Labeling mixtures (100 pl) contained DNA at 0.75 pg (for low resolution nuclease protection studies) or 0.125 pg (for high resolution nuclease protection studies) and 300 pCi y-(KP)- 37 ATP (7000 Ci/mmol; ICN). To generate DNA fragments labeled at their 3* ends, the appropriate restriction fragment containing a 5' overhanging end was filled in to form a blunt end with unlabeled dATP, dCTP, and dGTP and radiolabeled dTTP by treatment with Klenow fragment (BRL) according to the procedure of Ausubel et al. (1989). The reaction mixture contained 0.8 pg DNA and 80 pCi a-(MP)-dTTP (3000 Ci/mmol;

Amersham) in a total volume of 30 pl.

After end-labeling, reaction mixtures were passed through a spun column

(Sambrook et ai., 1989) containing Sephadex G-50 (Pharmacia Inc., Piscataway, NJ) to remove unincorporated labeled nucleotides and were stored at -20°C. The probes used for S1 mapping analyses were extracted further with phenol-chloroform and two successive extractions with chloroform and precipitated with sodium acetate and ethanol overnight at -20°C. The probes were recovered by centrifugation, washed, dissolved in

20 pl of DEPC (diethyl pyrocarbonate; Sigma)-treated distilled H20, and stored at -20°C.

Following extraction as described above, the end-labeied primer used for generating sequencing ladders was precipitated with 0.5 volumes of 7.5 M ammonium acetate and

2.5 volumes of ethanol overnight at -20°C. The primer was recovered by centrifugation as described above and dissolved in 20 pl of distilled H20. Radioactivity incorporated into each S1 probe was measured in a scintillation counter (Model LS-7500; Beckman

Instruments Inc., Palo Alto, CA). The 5' and 3’ end-labeled probes used for low resolution SI analysis contained 600,000 and 750,000 cpm/pl respectively; the probe used for high resolution S1 analysis contained 95,000 cpm/pl. 38 DNA fragments labeled by the random primer method (Prime-A-Gene; Promega)

served as probes for colony and Southern hybridizations. Labeling mixtures (50 pi)

contained 25 ng DNA and 50 pCi of a-(“ P)-dCTP (3000 Ci/mmol; Amersham).

IV. Hybridization analysis

A. Southern hybridization

Genomic DNA from S. griseus or S. coelicolor or phage DNA was digested with various restriction enzymes and resolved on 1% agarose gels. The restricted DNA was transferred to nitrocellulose membranes (BA 85; Schleicher and Schuell Inc., Keene, NH) by the procedure described by Sam brook et al. (1989) or nylon membranes (Zeta-Probe;

Bio-Rad) by capillary elution under alkaline conditions according to the protocol of the supplier. After transfer of DNA, the membranes were baked for 1.5 hr at 80°C. When

DNA fragments were used as probes, the membranes were prehybridized at 37°C for 1 hr and hybridized with the probe overnight at 42°C. The prehybridization and hybridization solutions contained 50% formamide (Sambrook et al., 1989). After hybridization the membrane was washed at increasing temperatures (5°C increments from 55° to 75°C) in 2X SSC (1X SSC is 0.15 M NaCI and 0.015 M [Na],citrate) and 0.1% sodium dodecyl sulfate (SDS), and then in 0.2X SSC and 0.1% SDS. The washed membranes were exposed to X-OMAT film (Eastman Kodak Co., Rochester, NY) in the presence of two intensifying screens at -70°C. The film was developed in an automatic

X-ray film processor (CWP 14-Plus, Fischer Industries Inc., Geneva, IL).

When oligonucleotides were used as probes, the membranes were prehybridized at 37°C for 1 hr and hybridized with probe overnight at 37°C. The prehybridization 39 solution contained 5X Denhardt's solution (Sambrook et al., 1989), 10 mM EDTA

(disodium salt), pH 7.5, and 0.5% SDS, and the hybridization solution contained 6X NET

(Sambrook et al., 1989), 5X Denhardt’s solution, 0.1% SDS, and 250 pg/ml denatured salmon sperm DNA. After hybridization the membrane was washed four times for 10 min each in 6X SSC and 0.05% sodium pyrophosphate at room temperature and then in the same solution at 42°C for 30 min. The washed membranes were exposed to X-OMAT film as described above.

B. Plaque hybridization

A mixture of 20-mer oligonucleotides (Fig. 5) generated on the basis of the N- terminal 7 amino acids of purified histidase from S. griseus (Wu et al., 1992) was end- labeled and used as the probe in plaque hybridizations to detect the histidase structural gene in a library of S. griseus genomic DNA. The genomic DNA library in AEMBL4 was constructed by K. E. Kendrick and contained Sau3M fragments of approximately 10-20 kb prepared as a partial digest of S. griseus genomic DNA. E. coli LE392 was grown in

3 ml TSB supplemented with 10 mM MgCI2 and 0.2% maltose. Phage propagation and in situ plaque hybridization on a nitrocellulose membrane (Schleicher and Schuell) were performed as described by Sambrook et al. (1989). The probe was used at a concentration of 9 pmoi/ml in the hybridizations. The membranes were prepared, washed, and subjected to autoradiography for 2 days as described above. Plaques which showed positive hybridization signals on duplicate membranes were chosen for further purification. The screening procedure was repeated until all plaques on a given plate showed a positive signal. Fig. 5. Degenerate oligonucleotide used for cloning hutH.

40 41

Generation of the oligonucleotide probe

1. N-terminal sequence of histidase purified from Streptomyces griseus:

M et-Asp-M et-H is-Th r-Val-Val-Val-G ly-Th r-Ser-G ly-

2. From this sequence, a mixture of oligonucleotides was prepared:

5 ' ATGGACATGCACAGAGTAGT 3 ’ T T C C G G T T

Figure 5 42

C. Northern hybridization

Northern hybridization was performed by the method of Sambrook et al. (1989).

The RNA preparation (approx. 2.5 pg) was fractionated on a denaturing agarose-

formaidehyde gel and transferred to a nitrocellulose membrane (Schleicher and Schuell)

by capillary elution. The RNA on the membrane was hybridized with the degenerate

oligonucleotide mixture designed from the histidase N-terminaJ amino acid sequence.

The probe was used at a concentration of 9 pmot/ml. The membranes were prepared,

washed, and exposed to Kodak X-OMAT film for 16 hr as described above.

D. Colony hybridization

Colony hybridization was performed to identify E coli TB1 transformants that

harbored the hutH gene from S. griseus SKK896, a Hut mutant, in pUC18. Genomic DNA

of SKK896 (16 pg) was digested with SamHI overnight. The restricted DNA was fractionated by agarose gel electrophoresis. A portion of the gel containing DNA

fragments of approximately 4 kb in length was excised and the DNA was purified by the

phenol'freeze-fracture method as described above. The DNA was dissolved in 20 pi

distilled H20. Approximately 0.4 pg of fragment was ligated into the SamHI site of dephosphorylated pUC18 (0.1 pg) in a 20 pi reaction volume, and the resulting plasmids were introduced into E coii TB1. White colonies on LB agar containing ampicillin and

X-gal were transferred to the same medium (100 colonies/plate). In situ hybridization of lysed bacterial colonies on nitrocellulose membranes (Schleicher and Schuell) was performed according to the method of Sambrook et al. (1989). A randomly primed DNA fragment containing hutH from the wild-type strain of S. griseus was used as a probe at a concentration of 0.6 ng/ml. The membranes were prepared, washed, and exposed to 43

Kodak X-OMAT film for 17 hr as described above. Colonies showing strong hybridization

signals were purified, and the correct insert in pUC18 was confirmed by analysis of

restriction digests.

V. Nucleotide sequence determination

The nucleotide sequence was determined by the dideoxy-chain termination

method of Sanger et al. (1977) using Sequenase version 2.0 (USB). Double-stranded plasmid DNA in pUC18 was first denatured by treatment with alkali and subsequently hybridized with the appropriate primer according to the manufacturer's instructions. The sequencing reactions were conducted according to the instructions of the manufacturer

(USB) with the following modifications. The inclusion of 10% dimethyl sulfoxide (Sigma) in the annealing reaction reduced the background substantially. To overcome sequence compressions, 7-deaza-dGTP was used instead of dGTP, and the termination reactions were incubated at 45°C rather than 37°C. In addition, Sequenase at 0.4 units per pi, as recommended by the supplier, was used in the reactions. Deoxycytidine-S’-a-^S)- thiotriphosphate (a-“ S-dCTP, >1000Ci/mmol; Amersham) was used to label the synthesized DNA fragments. The labeled DNA fragments were resolved by electrophoresis on a 6.0% denaturing polyacrylamide gel containing (per liter): 57 g acrylamide, 3.0 g bis-acrylamlde, 500 g urea, and 1X TBE buffer (10.8 g Tris-HCI, 0.93 g NajEDTA (disodium salt), and 5.5 g boric add per liter distilled HaO). The gel was run at a constant power of 60 watts on an S2 sequencing apparatus (BRL). After electrophoresis, the gel was soaked in a bath containing 10% acetic acid and 12% methanol for 20 min to remove the urea. The gel was dried and exposed to Kodak X-

OMAT film for 2-3 days at room temperature. 44

Analysis of the nucleic acid and amino acid sequences was conducted using a variety of the Wisconsin Genetics Computer Group (Madison, Wl) programs (Devereux et al., 1984). The Frame program (Bibb et al., 1964) as modified by G. Kleman and W.

R. Strohl was used for the identification of protein-coding sequences. PAUP (Swofford,

1990) was used to establish the phylogenetic tree of histidine ammonia-lyase and phenylalanine ammonia-lyase. The similarity of the amino acid sequence of ORF183 from pSAM2 was identified by using the Blast program (Altschul et al., 1990). RNA folding properties were analyzed by using the Mulfold program (Jaeger et al., 1989; Zuker, 1989).

VI. Bacterial transformations

Preparation of competent £. coll TB1 cells with CaCI2 and transformation with plasmid DNA were conducted as described by Sambrook et al. (1989). Streptomycete protoplasts were prepared essentially as described by Hopwood et al. (1985). Heat- activated, pregerminated spores of S. griseus were inoculated into 50 ml of TSB supplemented with 5 mM MgCI2< 1.0% glycine, and 5% PEG-8000. For preparation of protoplasts of S. griseus on a small scale, a sporulated colony was inoculated into 3 ml of TSB supplemented with 5 mM MgCI2 and 1% glycine in a test tube (18 x 150 mm) containing a glass bead. Subsequent manipulations were applied to cells in a 1.5 ml microcentrifuge tube. To prepare protoplasts of S. IMdans, a sporulated colony was inoculated into 50 ml of TSB supplemented with 5 mM MgCI2. Following an incubation of 6-8 hr to allow for germination of the spores, PEG-8000 (final concentration of 15%) was added to the culture. After overnight incubation, glycine (final concentration of 0.5%) was added because glycine seemed to inhibit growth of S. IMdans mycelia. Mycelia of

S. IMdans were harvested 6-8 hr after the addition of glycine, whereas mycelia of S. 45

griseus were harvested for protoplasting during late exponential growth (ASOOnm=8*10).

After treatment with lysozyme, mycelial fragments and protoplasts were separated by

filtration through a glass wool plug in a syringe. Protoplasts were stored in P-buffer at -

70°C at a concentration of 4 X 10B/ml.

Transformation of streptomycete protoplasts with plasmid DNA was carried out by the rapid scale procedure described by Hopwood et al. (1985). Protoplasts of S. griseus

were regenerated on SpMR agar; S. IMdans was plated on R2YE medium. After

incubation at 30°C for 20 hr (S. griseus) or 18 hr (5. IMdans), the regenerating colonies were overlaid with 1.0 ml distilled H20 containing thiostrepton.

To overcome the severe restriction system of S. griseus, plasmid DNA in

streptomycete replicons was first introduced into and screened in S. IMdans TK24 and then in S. griseus strain Rfs2. Subsequently, the plasmid DNA prepared from strain Rfs2 transformants was introduced into other strains of S. griseus. Protoplasts of strain Rfs2 were prepared fresh and used for transformation on the same day. To maximize the transformation efficiency, 0.5 pg of the plasmids prepared from S. IMdans was introduced into strain Rfs2. To facilitate the introduction of the large bifunctional vector pXE4 or its derivatives into S. griseus, plasmid DNA prepared from £. coli or S. IMdans was treated sequentially with Sssl and A/ul methylases (New England Biolabs). The methylated DNA was incubated at 65°C for 10 min to inactivate A/ul methylase, and 0.5 pg of the DNA was introduced into strain Rfs2. 46 Homologous transformation into both S. griseus and S. IMdans TK24 routinely

yielded 10° to 10T transformants per pg of plasmid DNA. Although special efforts were

taken to increase the transformation frequency of heterologous DNA in S. griseus , in most

cases fewer than 15 transformants were obtained from a single regeneration plate.

Plasmids isolated from these transformants were examined carefully by restriction

mapping to avoid selection of plasmids that had undergone rearrangements (Chen et al.,

1992; Yu and Chen, 1993).

VII. RNA techniques

Precautions had to be followed to prevent RNase contamination when working

with RNA. Glassware was autoclaved then baked at 250°C for 8 hr. Plasticware was

autoclaved twice in succession. The electrophoresis tanks and combs were immersed

in 50 mM NaOH for at least 1 hr before use. Distilled H20 was treated with 0.1% (final

concentration) DEPC (Sigma) overnight at 37°C and then autoclaved twice in succession.

Aqueous solutions were prepared using DEPC-treated distilled HsO and were autoclaved.

Reagents that could not be autoclaved (e.g., ethanol and isopropanol) were prepared fresh. All solutions and reagents were stored as aliquots at -20°C until use.

A. Run-off transcription

In two separate reactions the plasmid pKK501 was linearized by digestion with fcoRI and H/ndlll. The linearized plasmids were extracted and precipitated as described above. The plasmid DNA was recovered by centrifugation, washed with 70% ethanol, dried, and dissolved in distilled H20. Run-off transcripts were generated in vitro by incubation either with T7 RNA polymerase (in the case of EcoRI-digested pKK501) or SP6 47

RNA polymerase (in the case of H/ndlll-cut pKK501) according to the protocol of the manufacturer (Promega). After RNA synthesis, RQI DNase (2 units; Promega) was added to each reaction mixture to remove DNA. The reaction mixture was extracted with phenol-chloroform, precipitated with sodium acetate and ethanol, dissolved in 10 pi distilled H20, and stored at -20°C.

B. Isolation of RNA

Isolation of RNA from S. griseus cultures was carried out by the method of Kirby et al. (Kirby et al., 1967) with modifications by H. Baylis and C. Smith (personal communication). Cultures of S. griseus were grown in 50 ml of either complex medium containing L-histidine or glutamate minimal medium. Mycelia were collected during mid- exponential growth (A300r(T, of 2-4 in complex medium and 0.5-0.6 in glutamate minimal medium) by vacuum filtration on a 25 mm diameter, 0.45 pm pore size nitrocellulose membrane (Schleicher and Schuell) and washed with 100 ml distilled H20. The membrane was transferred to a 50 ml disposable polypropylene tube (Becton Dickinson and Co., Lincoln Park, NJ) containing 5 ml of ice-cold 2X modified Kirby mix (Hopwood et al., 1985) and 14 g of autoclaved and baked 3 mm diameter glass beads. The tube contents were vortexed vigorously for 1 min to lyse the cells and centrifuged at 10,000 x g for 10 min at 4°C. The aqueous phase was transferred to 5 ml of phenol-chloroform and centrifuged; the phenol-chloroform extraction procedure was repeated three times until no precipitate could be visualized at the interface. Nucleic acids were precipitated by the addition of 0.1 volume of 4.0 M sodium acetate, pH 6.0, and an equal volume of isopropanol and chilled for at least 2 hr at -20°C. The nucleic acids were collected by centrifugation at 10,000 x g for 15 min, washed with 70% ethanol, dried, and dissolved 46

in 400 til distilled H20. The nucleic acids were re-precipitated as described above and

dissolved in 180 pi distilled H20 and 20 pi of 10X DNase buffer (0.5 M Tris-HCI, pH 7.8

and 50 mM MgCt2). RNase-free DNase I (13.5 units; Boehringer) was added to the

solution. After incubation at 37°C for 30 min, the digestion was stopped by extraction

with phenol and chloroform, and RNA was precipitated with sodium acetate and

isopropanol. The RNA was recovered and dissolved in 100 pi distilled H20. Aliquots of

the RNA solution were used both to determine the concentration of RNA

spectrophotometrically and to verify RNA integrity by electrophoresis on a 1% agarose

gel. The RNA suspensions were stored at -20°C after the addition of 0.1 volume of 4.0

M sodium acetate, pH 6.0, and 1.0 volume of isopropanol.

C. S1 mapping and primer extension analyses

To determine the length of the hutH mRNA and whether hutH expression is

regulated in S. griseus, low resolution S1 mapping was performed according to the

method of Hopwood et al. (1985). The appropriate end-labeled DNA fragment (100,000

cpm) was co-precipitated with 50 pg of RNA. The pellet was washed, dried under vacuum for 10 min, and dissolved in 30 pi of hybridization solution. The nucleic acids were denatured at 85°C for 30 min, and the temperature was then decreased slowly (over a period of 30-40 min) to an annealing temperature of 62°C. After overnight incubation

(15-17 hr) at 62°C, the solution was treated with 0.3 ml of S1 digestion buffer containing

150 units of S1 nuclease (BRL). S1 termination solution was added and S1 -protected

DNA-RNA hybrids were precipitated. The pellet was disolved in 10 pi alkaline loading dye, and 4 pi of sample was fractionated on a 1% agarose gel under alkaline conditions

(Hopwood et al., 1985). The gel was dried, covered with plastic film, and subjected to 49 autoradiography with an intensifying screen and Kodak X-OMAT film for 3 days at -70°C.

Hybridization blots were quantified by scanning the autoradiographs in a GS 300

scanning densitometer (Hoefer Scientific Instruments, San Francisco, CA).

High resolution S1 nuclease analysis was performed to identify the hutH

transcription start site in S. griseus. The procedures and conditions used to obtain S1 •

protected DNA-RNA hybrids were identical to those used in low resolution S1 mapping.

The hybrids were resolved on a 6% gel and compared to nucleotide sequence ladders

generated by using an end-labeled primer. This primer, complementary to nucleotides

88 through 112 of the hutH coding region, had the same 5' endpoint (nt 112) as the

probe. Nucleotide sequence ladders were prepared by the method described above for

DNA sequence determination except that a-^S-dCTP was replaced by nonradioactive

dCTP. After electrophoresis, the gel was dried and exposed to Kodak X-OMAT film for

3 days at -70“C.

Primer extension analysis was performed to identify the hutH transcription site by

using the AMV reverse transcriptase primer extension system (Promega) with some

modifications (Sambrook et al., 1989). The end-labeled 25-mer oligonucleotide (12.5

pmd) used for generating nucleotide sequence ladders in high resolution S1 nuclease studies was co-precipitated with 20 pg of RNA. The pellet was washed, dried, and dissolved in 6 pi of distilled HzO. After addition of 5pl of AMV PE 2X buffer (Promega), the nucleic acids were denatured at 80°C for 20 min, and the temperature was then decreased slowly (over a 30 min period) to an annealing temperature of 42°C. After incubation for 3 hr at 42°C, the reaction mixture (9 pi) containing the extension mix 50

(including AMV reverse transcriptase; Promega), actinomycin D (50 |ig/ml; Sigma), and

RNasin (1 u/pl; Promega) was added to the annealing solution. The primer extension

reaction was carried out at 42°C for 60 min. The primer extension products were

precipitated with 0.5 volume of 7.5 M ammonium acetate and 2.5 volume of 100% ethanol

and kept at -20°C overnight. The primer extension products were recovered by

centrifugation, washed, dried, and dissolved in 20 pi of loading dye (Promega). Samples

(6 pi) were heated at 90°C for 10 min and resolved by electrophoresis as described above. The gel was exposed to Kodak X-OMAT film for 2 days.

VIII. Preparation of cell extracts and enzyme assays

The preparation of crude cell extracts and the spectrophotometric methods for analysis of histidase and urocanase activities were carried out as described previously

(Kendrick and Wheelis, 1982; Kroening and Kendrick, 1987). The enzyme assays were conducted in a Kontron Uvikon 930 thermoregulated recording spectrophotometer

(Kontron Instruments Inc., Everett, MA). For histidase activity, absorbance was monitored at 277nm. The extinction coefficient for the product, urocanate, at this wavelength is 18.8 x 103 1 mol'1 cm'1 (Kendrick and Wheelis, 1982). For urocanase activity, the decrease in absorbance was monitored at 300nm, at which wavelength the extinction coefficient is

7.05 x 103 1 m o l1 cm 1 (Kendrick and Wheelis, 1982).

The preparation of crude cell extracts for catechol dioxygenase assay was performed according to the method of Ingram et al. (1989) except that mycelia were disrupted by high pressure as described previously (Kroening and Kendrick, 1987). 51

Catechol dioxygenase activity was determined spectrophotometrically by monitoring the rate of appearance of 2-hydroxymuconic acid semialdehyde (Zukowski et al., 1963). The absorption coefficient is 33 x 103 1 mol'1 cm 1 (Bayly et al., 1966). CHAPTER Itl

RESULTS

I. Cloning the hutH gene from S. griseus.

Southern analysis using a degenerate oligonucleotide probe (Fig. 5) derived from the N-terminal 7 amino acids of purified histidase from 5. griseus (Wu et al., 1992) was carried out to determine whether the oligonucleotide probe could be employed to screen a genomic library prepared from S. griseus SKK821 (wild-type) DNA. The results (Fig.

6) showed that genomic DNA from SKK821 contained a 4.0 kilobase (kb) BamHI fragment, a 7.6 kb M/ul fragment, a 0.9 kb NcoI fragment, and a 1.5 kb SalI fragment that hybridized with the probe. On the basis of these results, the oligonucleotide mixture was used in plaque hybridizations to detect the 5’ terminus of the histidase structural gene in a >4EMBL4 library of SKK821 genomic DNA (Fig. 7). Three recombinant phages (of 45,000 examined) which hybridized with the probe were identified. Two of them were identical, and all three contained overlapping DNA fragments with a 16 kb Sau3M fragment in common. These three recombinant phages were designated >tKK500 (the identical two) and >4KK501. The N-terminus of the hutH structural gene was localized by

Southern analysis to an 8 kb EcofW/BglW fragment in >IKK500 and a 2.5 kb EcoRI/BamHI fragment in >4KK501 (Fig. 8). Subsequently these two fragments were cloned separately into the EcoRI/BamHI sites of pGEM-4Z. Restriction maps of the resulting plasmids pKK500 and pKK501 are shown in Fig. 9. A flowchart describing the overall strategy used to isolate and subclone hutH from S. griseus is presented in Fig. 10.

52 Fig. 6. Southern analysis of total genomic DNA from S. griseus SKK821 probed

with oligonucleotides specific to the N-terminus of hutH. Total genomic

DNA from SKK821 was digested with BamH\ (lane 1), Mlu\ (lane 2), Ncol

(lane 3), or Sa/I (lane 4) and resolved on an agarose gel. Each lane

contained 4 pg of DNA. After transfer to a nylon membrane, the DNA was

hybridized with the end-labeled probe at a final concentration of 6.8

pmol/ml. The membrane was exposed for 72 hr. Molecular weight

standards (mw) are indicated in kilobases.

53 54

1 2 3 4 mw

< 6.13 < 3.50 < 1.84

Figure 6 Fig. 7. Identification of hutH from a genomic DNA library of 5. griseus SKK821.

The oligonucleotide probe was used in plaque hybridizations to detect the

5' terminus of the histidase structural gene in an SKKB21 genomic DNA

library. A and B: a plaque showing a positive reaction (indicated by an

arrowhead) in duplicate plates was selected for further purification. C: the

purified recombinant phage.

55 Figure 7 Fig. 8. Southern analysis of AKK500 and AKK501 with the oligonucleotide probe.

Above, restriction maps of AKK500 and AKK501. The boxes represent

approximately 16.5 kb of S. griseus DNA. The thin lines represent that

portion of DNA derived from the vector, AEMBL4. The regions of each

recombinant phage that hybridized to the radiolabeled oligonucleotide

mixture are indicated by shaded areas in the boxes. Abbreviations: B,

BaroHI; E, EcoRI; G, BgtU S, San. Below, the DNA from AKK500 (Lanes 1-

5) and AKK501 (Lanes 6-10) was digested with Bgl\\ (lanes 1 and 6), Bg/ll

and EcoRI (lanes 2 and 7), EcoRI (lanes 3 and 8), EcoRI and Sa/I (lanes

4 and 9), or Sa/I (lanes 5 and 10). Each lane contained 1 pg of DNA. The

DNA fragments were transferred to nitrocellulose membranes and probed

with the radiolabeled oligonucleotides at a final concentration of 2.5

pmol/ml. The membranes were exposed for 16 hr. Molecular weight

standards are indicated in kilobases.

57 58

2< . 3- k> b 1 . 2 k b

2 0 k b 1 6 J 5 _ k b ______^ t _ §.-2_ kb_ > > ♦ -

EB G G BSE M XKK500

10 kb

G G BS SE GGGG XKK501 u

10 kb

12345 6789 10

< 9 . 8 « < 5 .1 <^2.4 ^ 2 . 2

< 1 .3

Figure 8 Fig. 9. Restriction maps of pKK500 and pKK501. Thin lines and shaded boxes

represent vector DNA (pGEM-4Z) and S. griseus DNA, respectively. A

region in pKKSOO is shortened in the drawing and marked by double shills

(II). The arrow indicates the position of hutH and the direction of

transcription. Abbreviations : A, Sai/3AI; B, BamHI; D, Nde I; E, EcoRI; G,

Bg/ll; H, H/ndlll; M, M/ul; N, A/col; S, Sa/I; T, Ssfl

59 PKK500

pKKSOl Fig. 10. Strategy used to clone hutH from S. griseus SKK821.

61 62

S. griseus chromosomal DNA XEMB14

Sad3A\ partial digest V size fractionation \ # SamHI digest (10-20 kb) V

SKK821 Genomic Library

Plaque hybridization with oligonucleotide probe designed from histidase N-terminus

XKK500 UCK501

^ Southern hybridization ^

8 kb fcoRI-SgJI fragment 2.5 kb EcoRI-SamHI fragm ent

^ Ugation with FcoRI-SamHI-dlgested pGEM-4Z ^

pKKSOO pKKSOl I 3.45 kb EcoRI-H/ndlll fragment; Klenow, deoxyribonucleoslde triphosphates

I Ugation with Smal-dlgested pUC18 I pKK538 and pKK540 (two orientations)

Figure 10 63

Southern analyses of AKK500 and AKK501 (Fig. 8) and pKKSOO and pKK501

indicated that the portion of the hutH gene that encodes the N-terminus was located near

the EcoRI site in pKK501. Therefore it was unclear whether I had cloned the entire

histidase structural gene or only the upstream portion of hutH. Run-off transcription and

northern analysis were performed to determine the direction of transcription of the hutH

gene in pKK501 and to indicate whether or not the entire hutH structural gene had been

cloned in this plasmid. As shown in Fig. 5, the nucleotide sequence of the

oligonucleotide probe corresponds to the sense strand of hutH. If the hutH gene were transcribed from the EcoRI site toward the H/ndlll site (clockwise in Fig. 11), then only the transcript generated from the T7 promoter would hybridize with the oligonucleotide probe.

On the other hand, if the hutH gene were transcribed in the opposite direction

(counterclockwise in Fig. 11), then only the transcript generated from the SP6 promoter would be capable of hybridization with the oligonucleotide probe. The results (Fig. 11) showed that only a 2.5 kb transcript synthesized from the T7 promoter hybridized with the probe. On the basis of these experiments and the subunit molecular weight of purified histidase from S. griSBUS (Mr=55,000; Wu et al., 1992), the direction of the hutH gene in the 2.5 kb fragment in pKK501 is from the EcoRI site toward the H/ndlll site, and pKK500 is likely to contain the entire hutH gene.

II. Features of the nucleotide sequence of hutH

Because pKK501 might not have contained the entire hutH structural gene, I constructed pKK535 by replacement of the 0.67 kb M/ul/H/ndlll fragment in pKK501 with the 1.64 kb M/ul/H/ndlll from pKKSOO (Fig. 12). This replacement resulted in the reconstruction of the hutH structural gene joined to a greater extent of downstream Fig. 11. Determination of the direction of transcription of hutH in pKK501. Above:

schematic diagram of pKK501. The shaded region of pKK501 indicates

5. griseus DNA. The region of pKK501 that hybridized with the

oligonucleotide probe is marked by an asterisk. Below: in two separate

reactions, plasmid pKK501 was linearized by digestion with EcoRI or

H/ndlll, and T7 RNA polymerase or SP6 RNA polymerase was added to

obtain transcripts in Wfro. Transcription products were separated on a

denaturing agarose-formaldehyde gel, transferred to a nitrocellulose

membrane, and probed with the oligonucleotide mixture. Lanes 1 and 3

represent 2.6 kb and 8.0 kb molecular weight standards respectively; lane

2 contains the transcript made from the T7 promoter (EcoRI-cut template);

lane 4 contains transcript made from the SP6 promoter {W//id Ill-cut

template).

64 65

SP6 f EcdR\

1 2 3 4

# •

Figure 11 Fig. 12. Construction and restriction maps of pKK538 and pKK540. (A) The

shaded and unshaded regions of plasmids indicate S. griseus DNA and

vector sequences, respectively. The 0.67 kb M/ul/H/ndlll fragment in

pKK501 was replaced by the 1.64 kb M/ul/H/ndlll fragment from pKK500.

The resulting plasmid, pKK535, containing a 3.45 kb insert in pGEM-4Z,

was introduced into E. coli TB1. The ends of the 3.45 kb fcoRI/H/ndlll

fragment from pKK535 were made blunt with Klenow fragment and

inserted into the Smal site of p(JC18 to generate pKK538 and pKK540. (B)

Thin lines represent vector sequences; the shaded box indicates 5.

griseus DNA. Only the restriction map of pKK540 is shown; pKK538

contains the same insert in opposite orientation in pUC18. Abbreviations:

A, Sau3A; B. BamHI; D, A/del; E, EcoRI; G, Bgl II; H, H/ndlll; M, M/ul; N,

A/col; S, San-, T, Ssfl.

66 H B H

Cut with H/ndlll and M tuI Cut with H/ndlll and MliA

E M H H \ pKK535

Cut with EcoRI and H/ndlll

M B H Ends filled in with Klenow fragment; hutW ligated to the Smsl site of pUC18

pKK540 6 .15 kb A.

A S N T N TSMSN B NSD HQ i i , i i i i m i i i i E pKK540

hutH coding B. sequence

Figure 12 68 genomic DNA. For unidirectional digestion with exonuclease III, the resulting fragment

(3.45 kb) was excised from pKK535. After making the ends blunt, the fragment was ligated into the Sma\ site of pUC18 in two orientations to generate pKK538 and pKK540

(Fig. 12). Plasmids pKK501, pKK538, and pKK540 were used for generating nested deletions to determine the nucleotide sequence of the 3.45 kb £coRI/H/ndlll fragment in both strands. The 48 deletion clones as well as pKK501, pKK538, and pKK540 used for determining the nucleotide sequence of this fragment are depicted in Fig. 13, and the nucleotide sequence is shown in Fig. 14.

Two streptomycete open reading frames (ORFs; Fig. 15) were identified in this fragment by using the Frame computer program (Bibb et al., 1984) as modified by G.

Kleman and W. R. Strohl. Computer searches indicated that ORF1 shows significant similarity (33% identity; 52% similarity at the amino acid sequence level) to ORF183, a gene thought to be involved in the regulation of replication of an integrating element, pSAM2, from Streptomycesambofaciens (Fig. 16; Hagdgeet al., 1993, and unpublished).

The overall G+C content of ORF2 is 74%, characteristic of the high G+C content of a streptomycete gene (Hopwood and Kieser, 1990). The N-terminaJ 12 amino acids deduced from the nucleotide sequence of ORF2 correspond exactly to those of purified histidase (Wu et al., 1992). ORF2 would be expected to encode a polypeptide of 516 amino acid residues with a molecular weight of 53.5 kDa, which is in agreement with the estimated subunit molecular weight of 55 kDa for purified histidase (Wu et al., 1992). The deduced amino acid sequence of the ORF2 gene product is similar to sequences of other bacterial and mammalian histidases, especially in the central portion of the primary structure (Fig. 17). It appears therefore that ORF2 is the histidase structural gene. On Fig. 13. Strategy for the determination of the nucleotide sequence of the 3.45 kb

EcoRI/H/ndlll fragment. Arrows indicate the length and direction of the

nucleotide sequence determined in each exonuclease Ill-generated

subclone of pKK501, pKK538, and pKK540.

69 EcoRI Midi H/ndlll

0 500 1000 1500 2000 2500 3000 3444

< 3 = <3=

< 5= < 3 =

from pKKSOl from pKK538 from pKK540

Figure 13 Fig. 14. Nucleotide and deduced amino acid sequences of the 3.45 kb

EcoRI/Hind\\\ fragment. ORF1 indicates the partial open reading frame of

unknown function. Putative promoter regions are marked by asterisks.

Two hutH transcription start sites determined by S1 mapping (Fig. 23) are

designated "+1". A potential transcription terminator located immediately

downstream of hutH is indicated by arrows. Other inverted repeats are

marked by triangles. The double-underlined region designates A-like DNA.

The hutH sequence has been deposited in Gen Bank under the accession

number M77841.

71 IVLHGRA*GELYVARQAGQPVFDRADADFATVL GATCGTGCTGCACGGGCGGGCGTGGGGTGAGCTGTACGTGGCGCGGCAGGCCGGACAGCCCGTCTTCGACCGGGCCGACGCGGACTTCGCGACCGTCCTC • • • • • • • • • 100 . . .0R F1**+^

AAVVASGIAQTERLEEVRKLAFTOPLTGLANRRA GCCGCCGTCGTGGCCTCCGGGATCGCCCAGACCGAGCGCCTGGAGGAGGTCCGCAAGCTGGCCTTCACCGATCCGCTGACCGGTCTGGCGAACCGCCGGG • •••••••• 200

VDVRLDEAMERRRVDATVVSLVVCOLNGLKAVN CGGTCGACGTCGGGCTOGACGAGGCCATGGAfiCGCCACCGTGTCGACGCCACGGTCGTCAGCCTOGTCGTCTGCGATCTGAACGGCCTCAAGGCGGTGAA • • • • • • • • • 300

DTHGHAVGDRLLERFGSVLSLCGAMLPEALAAR CGACACCCATGGCCACGCCGTCGGCGACCGCCTGCTGGAACGTTTCGGCTCGGTGCTCTCCCTGTGCGGGGCGATGCTGCCCGAGGCGCTGGCGGCCCGG » • • • • • • • * 400

LGGOEFCLLTAGPPAOAVVGVATELCDRAAVIEL CT GGGCGGCGATGAGTT CT GCCT GOT GACGGCGGGGCCCCCGGCCGACGCGGTGGT CGGCGT CGCCACCGAGCT GTGCGACCGGGCGGCGGT GAT CGAGC * • • • • • • • • 500

GDGVACGVASTGDPIGPVRSARRLFRLADAAQY TGGGCGACGGGGTCGCCTGCGGGGTCGCCTCCACCGGCGACCCGATCGGCCCGGTGCGCTCGGCCCGGOGGCTGTTCCGGCTCGCGGACGCCGCCCAGTA • • • • • • • • • 600

RAKAARAPGPVVAGRDGEVVRLADSPPKSAHOR CCGGGCCAAGGCGGCCCGCGCGCCGGGCCCCGTGGTGGCCGGGCGGGACGGCGAGGTCGTCCGGCTCGCGGACTCCCCGCCGAAGTCCGCGCACGACCGG • • • • • ■ • • • 700

RRLRGNRP o n d 4444444444-44444444 * * CGCAGACT GCGCGGCAACCGGCCGT GAGCCGGGGT GCGCCCGT GCCGT CGCCGT CGCGGGGCGCCCCCT CCCGT AAGGGGCCCACCGCGGCACCACTAGT « • • • • • • • • 800

-35 -10 +1 *1 ...... ***• momhtvvvgtsgttaeovvava GACATGAAGGCTTTCACTAGGTACGCTGCTGAATATGGATATGCACACTGTTGTGGTGGGGACGTCCGGAACCACCGCCGAGGACGTCGTCGCCGTGGCC • • • • • • • • • 900 h u tH ( 0 R F 2 )- m ^

RHGARVELSAAAVEALAAAR L I VDALAAKPEPVY CGCCACGGCGCCCGGGT CQSGCT CT CCGCCGCCGCCGTGGAAGCCCT GGCCGCCGCCCGT CT CAT CGTGGACGCCCT CGCCGCCAAACCCGAGCCGG TCT • 1000

GVSTGFGALASRHIGTELRAQLQRN1VRSHAAG ACGGTGTCTCCACCGGCTTCGGCGCCCTCGCCAGCCGCCACATCGGCACGGAACTGCGGGCGCAGCTCCAGCGCAACATCGTCCGCTCGCACGCGGCOGG • ■ • • • • • • *1100

MGPRVEREVVRALMFLRLKTVASGHTGVRPEVA CATGGGCCCGCGCGT CGAGCGCGAGGT CGT CCGCGCGCTGATGTT CCT CCGGCTGAAGACGGT CGCCT CGGGGCACACCGGCGTACGCCCCGAGGTCGCG • •••••••• 1200

Figure 14 Figure 14, coot

QTNADVLNAGITPVVHEYGSLGCSGOLAPL SHCA CAGACCATGGCCGACGT GCT GAACGCGGGCAT CACGCCCGT CGT CCACGAAT ACGGCT CGCT CGGCT GCT CCGGCGACCT CGCCCCGCT CT CGCACT GCG • ■■•••••• 1300

REKEGLALLNGTDGNLGMLVMALA0LRNLYTSA CCGCGAGAAGGAGGGCCT GGCCCTCOT CAACGGCACCGACGGCAT GCTCGGCATGCTGGT CAT GGCCCT CGCCGACCT GAGGAACCT CTACACCT CGGCC • 500

OITAALSLEALLGTDKVLAPELHAIRPHPGQGVS GACATCACCGCGGCCCTGTCCCTGGAGGCGCTCCTCGGTACGGACAAGGTCCTCGCCCCCGAGCTGCACGCCATCCGCCCGCACCCCGGACAGGGCGTCA • •«•**••• 1600

1700

DGRVESNGNFHGAPVAYVLDFLAIVAADLGSICE GACGGACGCGTCOAGT CCAACGGGAACTT CCACGGGGCGCCCGTCGCGT ACGT CCTGGACTT CCT CGCGAT CGT CGCCGCCGACCT CGGCT CGAT CTGCG • •••••••• 1900

RRTDRLLOKNRSHGLPPFLADDAGVOSGLMIAQ AGCGCCGCACCGACCGGCTGCTCGACAAGAACCGCTCGCACGGCCTGCCGCCGTTCCTCGCGGACGACGCCGGGGTCGACTCGGGCCTGATGATCGCCCA • •••••••• 2000

YTQAALVSEMKRLAVPASADStPSSANQEDHVS GTACACCCAGGCCGCCCTGGTCAGCGAGATGAAGCGGCTCGCGGTCCCCGCCTCCGCCGACTCCATCCCGTCCTCCGCGATGCAGGAGGACCACGTCTCC • •••••••• 2100

MGWSAARKLRTAVDNLARIVAVELYAATRAIELR ATGGGAT GGT CCGCCGCCCGCAAACTCCGT ACGGCCGT GGACAACCT GGCCCGGAT CGT CGCCGT CGAGCTGTACGCGGCGACCCGCGCCAT CGAGCT CC • •••••••• 2200

AAEGLTPAPASEAVVAALRAAGAEGPGPORFLA GCGCCGCCGAGGGCCTCACCCCGGCCCCCGCCTCGGAGGCCGTCGTGGCCGCGCTGCGGGCCGCCGGCGCCGAGGGCCCGGGCCCGGACCGCTTCCTCGC • 2300

PDLAAADTFVREGRLVAAVEPVTGPLA end »*»»»»»»»»» GCOGGACCTGGCCGCCGCCGACACGTTCGTACGGGAGGGGCGCCTGGTCGCGGCCGTGGAGCCCGTCACGGGGCCGCTGGCCTGAGCCGGTCCCGGACCC • •••••••• 2400

GCGAAAGAGGCCCCGTGCCGTGGCGCGGGGCCTCTCGCGT ACGCACGGGACCGCTCAGAGCGCCGGGCGCGCCCCGGATCCGGAGCGGCGCACGGAGTAG • •••••••• 2500 Figure 14, cont

GTGACGAAGCCTGCGCCGAGGCCCAACAGGGCTGTGCCGCCGATGAGAT ACGGGGTCGTGT CCGGGCCGGGCCCGGTGT CCGCGAGGAGCGCGCCGGAGG • ••••»•** 2600

CGTCGT CGGGGCTCCCGGCGGGCCGTT CGGCGGCT CGGGCGGAC6CG6CGGCCGGGGCGCCCTGAT CGCCGTGCT CACCGGT CGOGTTGGCCGACGGGAC • •••#•*** 2700

GAACCAOAOAGCGGCCAGCAGCGT GCCTGCGGCGGT GGCGGT CAGCAGT GTGCGACGAGT GACGGACACAGATT COAT CCCCTT GCGACAACCAT GAGTT • ••••••*• 2600

• •••*• + •• 2900

TTT GTT CGACTT CGCGT GGAGAT GGTGCT GGAGATCCGT CGACGCAGAT CCGCCTACCTTTCACGAGTT GCGCAGTTT GTCTGCAAGACT CT ATGAGAAG • • • •

CAGAT AAGCGAT AAGTTT GCT CAACAT CTTCTCGGGCAT AAGT CGGACACCATGGCAT CACAGT AT CGT GATGACAGAGGCAGGGAGTGGGACAAAATTG i i i i i i i i i aiuu

AAATCAAAT AATGATTTT ATTTTGACT GATAGT GACCTGTT CGTTGCAACAAATTGAT AAGCAAT GCTTTTTTAT AAT GCCAACTTAGT AT AAAAAAGCT ■ ■ 11 11 i ■ 1 ■—r* i i i 1 1 'i i i . j^ u u ■iGAACGAGAAACGT , AAAAT GAT , ATAAAT AT CAAT, AT ATTAAATT , ..I AGATTTT , 6CAT..... AAAAAACAGACTACAT— r — | AATACTGT AAAACACAACATATGCAGT | | imuCAC

T AT GAAT CAACTACTT AGATGGT ATTAGT GACCT GT AACAGAGCATT AGCGCAAGGT GATTTTT GTCTT CTT GCGCT AATTTTTTGT CAT CAACCTGTCG ■ 1...... "T" I I I " 1 I I I I '"'"WOP

GCACT CCAGAGAA6CACAAAGCCTCGCAATCCAGTGCAAAGCTT 3444 Fig. 15. identification of protein-coding sequences in the 3.45 kb £coRI/H/ndlll

fragment by Frame analysis. Because streptomycetes contain DNA with

a high G+C content (73%), their protein-coding regions typically have

intermediate (60-80%), low (50-60%), and high (90-97%) G+C contents in

the first, second, and third positions of the codons, respectively (Bibb et

al., 1984). By this analysis, one can readily identify coding sequences.

75 100 90

500 1000 Nucleotide

■ Frame 1 0 Frame 2 ■ Frame 3

Figure 15 Fig. 16. Alignment of the deduced amino acid sequence of ORF1 with ORF183

from Streptomyces ambofaciens. \,identical amino acids; conservative

amino acid substitutions.

77 1 . . . IVLHGRAWGELYVARQAGQPVFDRADADFATVLAAWASGIAQTERL 47 O rf 1 1...... VDPQTIa T L LPLLGwll-HGSVL...... IrAl 26 pSanfi 0rf183

48 EEVRKLAFTDPLTGLANRRAVDVRLDEAMERHRVDATWSLWCDLNGLK 97 Orf1

27 AS a Ar AA l U l Ht Aa GWTa Aa EHCIH Ah PRa A VLL v A l DHFK 68 pSam2 Orf 183

98 AVND HGHAVGDRL L ER FGSVL. SLCGAMLPEALAARLGGOEFCLLTAGP 146 O r f1

69 TLIL i m m AALIATAHRLS ti U . ..rhgt Ag U m r r AIRDL 115 pSam2 Orf 183

147 PADAWGVATELCDRAAVIELGDGVACGVASTGOPIGPVRSARRLFRLAD 196 O rf1

116 DAVDLDALTtALHQPTNYDGTALPLAASVGVCRVAELPVPALTDALAAADJ iLpdLvSVGVCRVAELAvPALTDALAAiA 165 pSam2 Orf 183

197 AAQYRAKAARAPGPWAGRDGEWRLADSPPKSAHDRRRLRGNRP* 242 Orf1

166 UmyaAkG ...... A s A . AA sAA a R* 184 pSanfi Orf 183

Figure 16 Fig. 17. Alignment of histidase primary structures. The amino acid sequences of

histidases from Bacillus subtllis (Oda et al., 1988), Pseudomonas putida

(Consevage and Phillips, 1990), and rat (Taylor et al., 1990) are aligned

with that of S. griseus. Only amino acid differences are shown; dots

indicate identical amino acid residues, and gaps are marked by dashed

lines. The alignment was generated using the Wisconsin Genetics

Computer Group program PILEUP. Similarity indices of aligned histidase

tabulated with the percent similarity (numerator) and percent identity

(denominator) in pairwise comparisons are shown at the end of the

sequence alignment. These values were calculated using the Wisconsin

Genetics Computer Group program GAP with a gap penalty of 5 and a

gap length penalty of 0.3.

79 80 S. griseus P. putida B.~subtilis ~ Rat MPRYTVHVRG EWLAVPCQDG KLSVGWLGRE AVRRYMKNKP DNGGFTSVDE VRFLVRRCKG LGLLDNEDLL EVALEDNEFV 81 160 S. griseus ...... MD MHTVWGTSG TTAEDWAVA RHGARVELSA AAVEALAAAR LIVDALAAKP P. putida L.LAQ.R..H AAPV..Q.D. S.AP..O.SV AC..Q.I.ED B. s u b tilis L.TA..AR.L FDFEEAAA.E ES..R.KKS. AA..R.VRDE Rat EWIEGDVMS POFIPSQPEG VFLYSKYREP EKY.A.DGDS i .T .... N.* .GHY..K.TS I.EKK.QQS. E...S.IKER 161 240 S. griseus EPVYGVSTGF GALASRHIGT ELRAQLQRNI VRSHAAGMGP RVEREWRAL MFLRLKTVAS GHTGVRPEVA QTMADVLNAG P. putida RTA...N... • L k • « 1 ft « . . HDLE___S. ■ L...... A ...D....L. .V...N..SR .F___RK.I DA. IA___E B. s u b tilis KT*« t *N<.■ .KFSDVL.QK .OS.A..L.. •L...C...D •FPEC.S... .L..ANA.LK . F... . A. . I EQ.LAF..KR Rat TV .KF.RTV.PA NKLQE..V.. .R..SS... K . .SP.RC.M. .A.. .NV..K .Y.. .SL.TL KQ.I ..F. .S 241 320 S. griseus ITPWHEYGS LGCSGOLAPL SHCALTLMGE GEAEGPDGTV RPAGELLAAH GIAPVELREK EGLALLNGTD GMLGMLVMAL ~P. putida .Y.H.PLK.. ••A...... AT1IS.V___ .K.RY-K.QW LS.T.A..VA . .E..T.AA...... Q .ST.YA.RG. B. s u b tilis .H.. .PQQ.. ■•A...... L..A...Q . .VFF-E.ER M..MTG.KKA ..Q..T.TS...... Q ..T..G...Y Rat CLSY.P.K,. . .A...... L..G ______.KMWS.KSGW AD.KY..E.. . .K..V..P...... Q M.TS.GCE.. 321 400 S. griseus AOLRNLYTSA OITAALSLEA LLGTDKVLAP ELHAIRPHPG QGVSAONMSR VLAGSGLT-- -GHHQODAPR VQDAYSVRCA ~P. putida FYAED..AA. I AC...... ______RSPFDA R..EA.GQ.. .ID..ACFRD . ..---D.SE VSLSHKNCD. .. .P ...... Q B. subtilis I.AEK.AYQT kRIkSt■■■• .Q.IIDAFOE . . .LA.GYQE .IDV..R.RF Y.SD______TS Q.EL...... D Rat ERASA.ARQ. ..V ...... V ,K. .T.AFDT t t t t t t i * FI • . IEV.FRFRS . .DSDKHPSE I.ESHRFCD...... C 401 480 S. griseus PQVNGAGRDT LDHAALVAGR ELASSVDNPV VLPD-GRVES NGNFHGAPVA YVLDFLAIVA ADLGSICERR TDRLLDKNRS P. putida .. .M..CLTQ .RQ..E.L.I .ANAVS______.FAAE.D.I. G. . . .AE... MAA.N.. .Al ...... S... ISL....HM. B. s u b tilis ...H..T¥(Q. .GYVKEKLEI . .NAAT______.FN.GD..1. ^3. . . . . Q... .A.... K.Al S..AN.A... I ______NP.LN Rat ...H.WN.. .AFVKD.ITT .. N.AT.... .FASR.ETI. G...... EYP. KA...... GV H..AA.S... I . ..CNPSL.

Figure 17 Figure 17, cont 481 560 S. griseus HGLPPFLAOD AGVDSGLMIA QYTQAALVSE MKRLAVPASA DSIPSSAMQE DHVSMGWSAA RKLRTAVDNL ARIVAVELYA ~P. putida Q- V.N .V.A...A.. N.A.SH.H.V...... N...... APA...... WEMA. .T RG.P...WL. B."subtilis D- SPH P..Q..A..M . .AA.S N.T..H...V ...... N...... TI.. .HAYQV.A.T R AIC Rat E-..A..VA. HC.A S.A.CH.S.V ...S...AT...... GW.. ..ALRV..H. EQ L. 561 640 S. griseus ATRAIELRAA EGLTPAPASE AWAALRAAG AEGPGPDRFL APDLAAADTF VREGRLVAAV EPVTGPLA*...... P. putida .CQ -- K..KTSAKL. KARQ...SEV .-HYDR...F EK.VEL .AK.S.T.L. PAGVL.SL*...... B. s u b tilis .LQ...Y.-- GIEHA.SY.K Q.FQE..KW P-SIQQ..VF SY..ERLT0. ..KES..PDH QNKELRGHNI * ...... Rat .CQ...FL** RP.KTTTPL. K.YDL..SW R-PWIK...... E..HRL .L.Q..WEVA A.YIEKYRME HIPESRPLSP 641 662 S. griseus P. putida B. aubtHIa Rat S. griseus ...... 100/100 56/38 52/38 52/37 ~P. putida ...... 100/100 57/42 58/42 B. s u b tilis ...... 100/100 56/42 Rat TAFSLESLRK NSATIPESOD L* 100/100 82 the basis of the alignment of amino acid sequences of histidine ammonia-lyase and the analogous enzyme phenylalanine ammonia-lyase (EC 4.3.1.5), a phylogenetic tree was established to identify their lineages (Fig. 18). The tree indicated that histidase from S. griseus is related to phenylalanine ammonia-lyase, and that histidase from S. griseus is divergent from the other bacterial and mammalian histidases.

Inspection of the nucleotide sequence surrounding the hutH structural gene revealed several interesting features (Fig. 19). The regions located 10 nucleotides and

35 nucleotides upstream of the translation initiation codon resemble the consensus sequence of E. co//-like streptomycete promoters (Strohl, 1992). A stem-and-loop structure (AG=-42.9 Kcal/moi) immediately downstream of hutH resembles a factor- independent transcription terminator (Deng et al., 1987). The nucleotide sequence downstream of the hutH coding sequence (from nucleotide 2947 to nucleotide 3444) is identical to that of A DNA within the int gene (from nucleotide 27479 to nucleotide 27976;

Hendrix et al., 1983), which also contains the att site.

To determine whether S. griseus chromosomal DNA contains A DNA downstream of hutH, Southern analyses were performed. A 1.87 kb EcoRI/SamHI fragment was isolated from A DNA which spanned the int gene and the adjacent upstream region of the

A genome (nt 26104 to 27972). This fragment did not hybridize to genomic DNA from S. griseus SKK821 (data not shown). In contrast, intense hybridization signals were observed when a 0.51 kb EcoR\/Sal\ fragment containing the entire A-like DNA plus an additional 3 bp of 5. griseus DNA from pKK540 was used as a probe (Fig. 20, lanes 2-4). Fig. 18. Dendrogram of the relatedness of amino acid sequences of histidine

ammonia-lyases and phenylalanine ammonia-lyases from plants and fungi.

The tree was constructed by the computer program PAUP, which

incorporates maximum parsimony analysis. Sequences of the proteins

were obtained from the SwissProtein database.

83 P. putida Rat

B. subtilis S. griseus

plant and fungal phenylalanine ammonia-lyase (47% similar; 30% identical)

Figure 18 s Fig. 19. Features of the nucleotide sequence of the 3.45 kb EcoRI/H/ndlll fragment.

Double shills (//) indicate a region that has been shortened in the drawing.

Consensus sequences for E. co//-like streptomycete promoters (Strohl,

1992) are indicated below the putative hutH promoter. Two stem-and-loop

structures located upstream and downstream of hutH are indicated.

Transcription initiation sites suggested by S1 mapping are marked with

+ 1.

85 hutH

-36? -10? +1

GCACCACTAjsTGAC^naAAGGCTTTCACTACG rACGCTjjCTGAATATGGATATGCACACTGTTGTGGTGGGG ttg aca tagggt MDMHTVVVG g caa c

Figure 19 Fig. 20. Southern hybridization of total genomic DNA from S. griseus SKK821 with

X DNA. Only relevant restriction sites in pKK540 are shown. The positions

of nucleotides refer to those indicated in Fig. 14. Above, DNA fragments

used as probes. Regions of hutH and A-like DNA as well as three probes

(A, B, and C) derived from pKK540 are indicated. Abbreviations: B,

BamHI; E, EcoRI; H, H/ndlll; M, Mlu\, N, A/col; S, Sal]. Below, total

genomic DNA from SKK821 was digested with Sa/I (Lanes 2 and 5), Sa/I

and H/ndlll (Lane 3), flamHI (Lanes 4 and 7), or Sa/I and SamHI (Lane 6).

Each lane contained 3 pg of DNA. The restricted DNA was resolved by

electrophoresis in agarose gels, transferred to nylon membranes, and

hybridized with probe A (Lanes 2-4) or probe B (lanes 5-7) at final

concentrations of 1.25 ng/ml. The membranes were exposed for 24 hr.

Molecular weight standards (Lane 1) are indicated in kllobases.

87 88

H SS SMS B S E 203 242 1777 1606 1975 2476 2936 3444

PKK540 m

hutH coding sequence Probe A: 0.51 kb Probe B: B 0.98 kb Probe C: 3.45 kb

1 2 3 4 5 6 7

6.13 ? 3.50

-r.\‘

0.96

Figuro 20 89

Thus, it was not clear from these hybridization reactions whether the A-like DNA originated from S. griseus genomic DNA or was a cloning artefact. Because of the possibility that

DNA rearrangement occurred during construction of this cloned fragment, it was enticed to know the extent, if any, of rearrangement in pKK540. Two probes derived from

pKK540 were hybridized with SKK821 chromosomal DNA. One 0.98 kb EcoRi/SamHI

probe contained the entire A-like DNA and adjacent upstream DNA (Fig. 20, lanes 5-7); the other was the entire 3.45 kb £coR!-H/ndlll fragment (Figs. 20 and 21). The results indicated that the molecular sizes of the hybridized DNA fragments were as predicted from the restriction map from the HindUl site to the rightmost Sa/I site in pKK540.

Therefore the region of pKK540 extending from nt 1 to nt 2938 (the rightmost Sa/I site of

Fig. 20) showed no evidence of DNA rearrangement.

Southern analysis of total genomic DNA from S. griseus and S. coelicolor by using hutH from pKK540 as probe indicated that S. griseus appears to contain only one copy of hutH, and there is significant sequence similarity between S. griseus hutH DNA and S. coelicolor DNA (Fig. 21).

III. Complementation analysis

Complementation analysis was performed not only to confirm that I had cloned a functional hutH gene, but also to determine whether a S. griseus mutant (SKK896) deficient in histidase activity (Kroening and Kendrick, 1987) is defective in the histidase structural gene. To construct pKK1100, a 2.5 kb BamHI fragment containing the entire

ORF2 (the putative hutH gene) and the adjacent 0.83 kb of upstream DNA from pKK540 was ligated into the Bg/ll site of a multi-copy vector, plJ702, and introduced into SKK896 Fig. 21. Southern analysis of total genomic DNA from S. griseus and S. coelicolor

with the hutH structural gene as probe. Total genomic DNA from S.

griseus SKK821 (lanes 1*4) and S. c oelicoior M145 (lanes 5-8) was

digested with Sa/I (lanes 1 and 5), Mlu\ (lanes 2 and 6), BamHI (lanes 3

and 7), or A/col (lanes 4 and 8), resolved by electrophoresis, and

transferred to nylon membranes. Each lane contained 3 pg of DNA. The

membranes were probed with a 3.45 kb EcoRI/H/ndlll fragment containing

hutH from pKK540 at final concentrations of 1.25 ng/ml and 2.5 ng/ml for

SKK821 and M145, respectively. The membranes containing DNA from

SKK821 were washed at a stringency that would retain hybrids in excess

of 90% identity, and those containing DNA from M145 were washed at a

stringency that would retain hybrids in excess of 65% identity. The

membranes were exposed to X-ray film for 5 hr and 15.5 hr respectively.

Molecular weight standards (S1 and S2) are indicated in kilobases.

90 91

1234 S1S2 5678

26.3 ► < 26.3 6.13 ► < 3.18 3.18 ► < 1.54 1.54 ► < 0.96 0.96 ► t < 0.22 0.22 >>

Figure 21 92

{hut-2) and SKK821 (wild-type). The putative hutH gene was inserted in the opposite orientation relative to the tyrosinase-encoding me/ gene in the vector so that ORF2 would not be transcribed from the met promoter. The SKK896 transformants containing ORF2 were able to utilize L-histidine as sole nitrogen source and showed a high level of histidase activity (Table 4), whereas transformants containing the vector alone remained

Hut*. Surprisingly, when transformants of both SKK821 and SKK896 containing ORF2 were grown in the absence of L-histidine, SKK896 transformants produced a much higher level of histidase than did SKK821 transformants. In other words, the SKKB96 transformant showed a high level of constitutive production of histidase. This result suggests that SKK896 may be a regulatory mutant. On the basis of the results of these complementation experiments and the deduced amino acid sequence of the cloned DNA, the entire hutH gene appears to have been cloned.

IV. Repressor titration experiments

When pKK1100 was introduced into the wild-type strain, transformants contained significant levels of both histidase and urocanase activities when grown in the absence of L-histidine (Table 4). This result suggested that multiple copies of the hutH gene or its upstream sequence could sequester a negative regulatory factor. To determine whether expression of hutH is subject to negative control, repressor titration experiments were performed. Three fragments containing 67,176, or 333 nt upstream and 88, 88, or

78 nt downstream, respectively, of the hutH initiation codon were inserted into plJ702.

The copy number of p!J702 in S. griseus is approximately 50 per chromosome (K.E.

Kendrick, unpublished). The resulting plasmids were introduced into SKKB21. Although 93

Table 4. Hut enzyme activities in transformants of SKK821 and SKK896.

Specific Activity (nmol/min/mg protein)*

Plasmid Host + Histidine" - Histidine" Strain Histidase Urocanase Histidase Urocanase

none SKK896 <0.1 <0.1 <0.1 <0.1 plJ702 SKK896 <0.1 <0.1 <0.1 <0.1 pKK1100 SKK896 74.04 ± 3.66 4.48 ± 2.27 80.56 ± 4.27 1.07 ± 0.24 plJ702 SKK821 8.39 ± 0.76 4.36 ± 0.32 <0.1 <0.1 pKK1100 SKK821 62.97 ± 2.75 4.37 ± 0.40 13.00 ± 1.85 0.46 ± 0.05

* Values are the average ± std. dev. of enzyme activities in at least two extracts, each assayed in triplicate. b + Histidine, CAA medium containing 10 mM L-histidine; - Histidine, glutamate minimal medium 94 the levels of histidase and urocanase were constant in all three transformants grown under inducing conditions, the transformants produced higher histidase and urocanase activities than did SKK821 containing only the vector when grown in the absence of L- histidine (Table 5). These results suggested that expression of both hutH and hutU is under negative control, and DNA residing between 67 and 176 nt upstream of the hutH coding sequence is required for this effect.

V. Transcriptional analysis of the hutH gene

Analysis of the nucleotide sequence revealed an E. coli-like streptomycete promoter consensus sequence preceding the hutH initiation codon (Figs. 14 and 19).

In addition, the complementation studies suggested that a promoter was located within the 835 nucleotides upstream of the hutH initiation codon (Table 4), although I cannot exclude the possibility of read-through from a promoter residing on the vector. To determine the existence and boundaries of a promoter-active fragment upstream of the histidase structural gene, a 150, 265, or 380 bp restriction fragment containing 67,176, or 291 nt, respectively, of DNA upstream of the hutH coding sequence and adjacent downstream DNA was inserted into the promoter-probe vectors plJ486 and plJ487 (Ward et al., 1986), such that the fragment was inserted in both orientations relative to the neo reporter gene that confers resistance to kanamycin (Fig. 22). The recombinant plasmids were introduced into S. lividans TK24 and 5. griseus SKK821. Kanamycin resistance was measured by assessing the extent of growth on minimal agar medium containing either

L-glutamate as sole carbon and nitrogen source or L-glutamate plus L-histidine. The results (Table 6) show that although the levels of kanamycin resistance in the S. griseus Table 5. Titration of a negative regulatory factor by upstream sequences in SKK821 and SKK896.

Specific Activity (nmol/min/mg protein)* Extent of • Histidine* Plasmid Upstream Host + Histidine* DNA (nt) Histidase Urocanase Histidase Urocanase

plJ702 — SKK821 8.39 ± 0.76 4.36 ± 0.32 <0.1 <0.1 pKK1101 67 SKK821 6.04 ± 0.49 4.52 ± 0.66 0.44 ± 0.21 0.15 ± 0.06 pKK1102 176 SKK821 9.28 ± 0.10 3.15 ± 0.26 1.17 ± 0.19 0.55 ± 0.07 pKK1103 333 SKKB21 7.48 ± 0.61 4.12 ± 0.50 1 39 ± 0.25 0.66 ± 0.12

pU702 — SKK896 <0.1 <0.1 <0.1 <0.1 pKK1101 67 SKK896 <0.1 0.65 ± 0.18 <0.1 0.51 ± 0.20 pKK1103 333 SKK896 <0.1 0.58 ± 0.08 <0.1 0.95 ± 0.31

Values are the average ± std. dev. of enzyme activities of at least two extracts, each assayed in triplicate.

+ Histidine, CAA medium containing 10 mM L-histidine; • Histidine, glutamate minimal medium.

Ui< 0 Fig. 22. Boundaries of DNA fragments used in promoter-probe studies. Putative

promoter-containing fragments (open boxes) and reporter genes (shaded

boxes) are diagrammed. Arrows represent the direction of transcription.

The ATG designates the translation initiation codon of hutH. The

nucleotides marking the endpoints of the promoter-containing fragments

are numbered as in Fig. 14.

96 Extent of Promoter Fragment Plasmid Vector97

ATG pKK1104 plJ486 768 835 923 060

pKK1105 plJ487 923 835 768 neo

PKK1106 plJ486 659 835 923 neo

pKK1107 plJ487 923 835 659 neo

PKK1108 plJ486 923 835 neo

PKK1109 plJ487 544 835 923 neo

pKK1112 pXE4 502 835 913

PKK1113 pXE4 913 835

pKK1114 pXE4 659 835 923 xyiE

pKK1115 pXE4 768 835 923 xyiE

Figure 22 Table 6. Promoter activity of the hutH upstream sequences using promoter-probe vectors plJ486 and plJ487.

Extent of Kanamycin Resistance (pg/ml)' Upstream Plasmid Sequence Glutamate* Glutamate + Histidine1* Orientation (nt) S. Hvidmns S. grismut S. IMctom S. griseus

plJ486 0 <1 0.1 <1 0.2 vector only plJ487 0 2 0.1 2 0.2 vector only pKK1104 67 5 0.1 5 0.2 correct pKK1105 67 <1 0.1 <1 0.2 incorrect pKK1106 176 5 0.1 10 0.5 correct pKK1107 176 <1 0.1 <1 0.2 incorrect pKK1108 291 <1 0.1 <1 0.2 incorrect pKKl 109 291 30 0.5 100 0.8 correct

* Maximum concentration of kanamycin sulfate that promoted normal growth on the indicated minimal medium. b Glutamate, minimal agar medium containing L-glutamate as sole source of carbon and nitrogen; glutamate + histidine, minimal agar medium containing glutamate and 10 mM L-histidine.

and 5. IMdans transformants are different, the pattern of resistance is similar in both

species. A hutH promoter appears to be located within 67 nt upstream of the hutH

coding sequence. Although the presence of DNA further upstream of hutH appears to

increase promoter activity in 5. IMdans, it is not clear whether L-histidine mediates this

effect.

To assess the strength of the hutH promoter more quantitatively, the promoter- probe vector pXE4 (Ingram et al., 1989) was used. This plasmid, containing a promoterless xyiE gene, is maintained at low copy number. The xyiE gene encodes catechol 2,3-dioxygenase, which oxidizes catechol, which is colorless, to a yellow product, 2-hydroxymuconic acid semialdehyde. The enzymatic assay is highly sensitive and readily monitored spectrophotometrically. A 156, 265, or 412 bp restriction fragment containing 67, 176, or 333 nt, respectively, of DNA upstream of the hutH coding sequence was inserted into pXE4 (Fig. 22) and introduced into SKK821. All fragments were inserted in the orientation that would permit transcription of xy/£, but only the 412 bp fragment was also inserted in the opposite orientation. The levels of catechol dioxygenase activity in extracts prepared from cells grown in the presence or absence of L-histidine (Table 7) indicated that the 67 bp fragment contains promoter activity, in agreement with the experiment using the promoter-probes pi J486 and plJ487. Extension of the upstream DNA in the pXE4 constructions did not significantly enhance XyiE activity.

The nucleotide sequence of the hutH region (Fig. 14) as well as promoter-probe studies suggested that either hutH is monocistronic or it is the first gene in an operon.

To determine the length of the hutH mRNA and whether hutH expression is regulated, Table 7. Promoter activity of 5. griseus transformants containing the hutH upstream sequences in the promoter-probe vector pXE4.

Catechol Dioxygenase (nmol/m In/mg protein)* Extent of Upstream Plasmid Sequences (bp) - Histidine * + Histidine11 Orientation

pXE4 — 0.26 ± 0.02 n.d.c vector only pKK1115 67 0.67 ± 0.02 3.72 ± 0.48 correct pKK1114 176 0.78 ± 0.05 4.93 ± 0.16 correct pKK1113 333 0.54 ± 0.03 0.66 ± 0.09 incorrect pKK1112 333 0.47 ± 0.05 5.16 ± 0.24 correct

Values are the average ± std. dev. for the enzyme activity from at least one extract assayed in triplicate.

- Histidine, minimal medium containing L-glutamate as sole carbon and nitrogen source; + Histidine, CAA medium containing 10 mM L-histidine. c n.d., not determined. 101

low-resolution S1 mapping was performed. Total RNA isolated from cultures of S. griseus

SKK821 grown in the presence or absence of L-histidine was hybridized with two probes

(Fig. 23): one (probe 1) was used to identify the approximate 5' end of the hutH

transcript, and the other (probe 2) was used to identify the approximate 3' end. Probe

1, which extended from nt 360 (approx.) to nt 1806, identified a 1.0 kb protected

fragment. Probe 2, which spanned nt 1806 to nt 2839 (approx.), identified a 0.6 kb

protected fragment. Thus, the total length of the hutH transcript is 1.6 kb. Although the same transcripts were evident in RNA isolated from cultures grown in the presence and absence of L-histidine, a ten-fold increase in band intensity was observed when RNA was

isolated from the culture grown in the presence of L-histidine.

The absence of an apparent ribosome binding site directly upstream of the translation initiation codon led to the possibility that transcription and translation of hutH may initiate at the same nucleotide position. High-resolution S1 nuclease analysis was performed to identify the hutH transcription start site. In the presence of L-histidine, the hutH gene has two apparent transcription start sites that occur at the first and seventh nucleotides of the hutH coding sequence (Fig. 24). Although the background was high, the results of primer extension analysis (data not shown) were identical to those of high- resolution SI nuclease studies. Transcripts initiating at the upstream site are 20-fold more abundant than those initiating at the downstream site. These results confirmed that transcription of hutH in S. griseus initiates at the nucleotide that marks the translation initiation site. In the absence of L-histidine, identical transcription start sites were aJso identified but at a 10-fold lower intensity. Fig. 23. Determination of the length of hutH mRNA. Above: schematic diagram

of the two probes. A 1.46 kb Hind\\\IMIu\ fragment (probe 1) from pKK570,

containing 1.0 kb of the hutH N-terminaJ coding sequence and adjacent

upstream DNA, was labeled at the 5’ end. Probe 2 was a 1.05 kb

Hind\\i/Mlu\ fragment derived from pKK546, which spanned 0.6 kb of the

hutH C-terminal coding sequence and adjacent downstream DNA. Probe

2 was labeled at the 3’ end. The labeled ends are marked by asterisks.

The other end (the H/ndlll site in both cases) was derived from the vector

pUC18, and the labeled end was therefore hydrolyzed by S1 nuclease.

Nucleotide sequence numbers refer to those indicated in Fig. 14. Each

probe (100,000 cpm) was hybridized to RNA (50 pg) prepared from

cultures of S. griseus SKK821 grown in the presence or absence of

L-histidine as indicated. tRNA (50 pg) from E. coli was used as the

negative control. Molecular weight markers are shown in kilobases.

Upper photo, using probe 1: lane 1, probe 1; lane 2, molecular weight

markers (kb); lane 3, RNA isolated from cells grown in the presence of

L-histidine; lane 4, negative control; lane 5, RNA isolated from cells grown

in the absence of L-histidine. Lower photo, using probe 2: lane 1, probe

1; lane 2, probe 2; lane 3, molecular weight markers (kb); lane 4, RNA

isolated from cells grown in the presence of L-histidine; lane 5, RNA

isolated from cells grown in the absence of L-histidine.

102 103 (835) ( 1806 ) (2385)

ATG TGAi— ~ l Mlu\

—* Probe 1: 1.46 kb

Probe 2: 1.05 kb

2.88 ►

1.54 ►

0.96 ►

0.52 ►

2.88 ►

1.54 ►

0.96 ►

0.52 ►

1 2 3 4 5 Figure 23 Fig. 24. Identification of the hutH transcription start site. A 0.6 kb BstM/Hind\\\

DNA fragment, containing 112 nt of the hutH N-terminal coding sequence

and approximately 475 nt of adjacent upstream DNA, was isolated from

pKK570 and labeled at the 5’ end (designated by an asterisk). The

H/ndlll-cut end of the probe was derived from pUCt 8. Lanes 1, 4, and 5,

RNA isolated from SKK821; Lane 2, RNA isolated from SKK896. The

probe (100,000 cpm) was hybridized to RNA (50 pg) prepared from

cultures of S. griseus grown in the presence (lanes 1, 2, and 4) or

absence (lane 5) of L-histidine. tRNA (50 pg) from E. coli was used as the

negative control (lanes 3 and 6). The sequence ladders (GCAT) were

generated by using an end-labeled 25-mer oligonucleotide that is

complementary to nt 88 through 112 of the hutH coding sequence. The

sequence ladders show the sequence of the antisense strand and are

written as the sense strand to the left of the autoradiogram. The

transcription start sites are marked by asterisks, and nt numbers are

designated as in Fig. 14.

104 105

GCAT 1 2 3GCAT 4 56

G G A T A* (841)

(835) (946)

fis/NI (probe: 0.6 kb) |-# (primer: 25-mer)

F igure 24 106

VI. Molecular characterization of 5. griseus SKK896, a Hut mutant

S. griseus strain SKK896 cannot utilize L-histidine as sole nitrogen source.

Enzymological studies (Kroening and Kendrick, 1989) and complementation analysis

(Table 4) of this mutant suggested that SKK896 may be defective in either the histidase

structural gene or the regulated expression of hutH. If the former possibility were the

case, then multiple copies of the hutH gene from SKK896 would not be expected to

suppress the Hut phenotype of SKK896. Southern analysis of total genomic DNA from

SKK896 and SKK821 with hutH from SKK821 as a probe was performed in order to confirm the presence of hutH in SKK896. The results showed that there is no visible difference in sizes of the hybridizing bands in the two strains (data not shown). On the basis of these experiments, the restriction map of hutH (Fig. 9), and Southern analysis using an oligonucleotide probe specific to the N-terminus of hutH with total genomic DNA of SKKB21 (Fig. 6), the hutH gene in SKK696 would be expected to lie in a 4.0 kb BamHI fragment. Subsequently, a 4.0 kb BamHI fragment containing the entire hutH gene and adjacent upstream sequences from SKK896 was cloned by colony hybridization with the

3.45 kb EcoRI-H/ndlli fragment containing hutH from pKK540 as a probe. This fragment was inserted into the Bgl II site of plJ702 to generate pKK1111 and then introduced into

SKKB21 and SKK896. The hutH gene from SKK896 was inserted in the opposite orientation relative to the mel gene in plJ702 so that hutH would be transcribed from its own promoter. SKK896 transformants containing hutH from SKK896 in a high copy number vector showed substantial histidase activity, even when grown in the absence of

L-histidine (Table 8). These results indicated that the histidase structural gene in SKK896 is at least partially functional. SKK896 is therefore more likely to have a regulatory defect. Table 8. Suppression of the defect in SKK896 at high copy number.

SpecHIc Activity (nmd/min/mg protein)

Plasmid Host Strain + Histidine* - Histidine* Histidase Urocanase Histidase Urocanase

pU702 SKK821 8.39 ± 0.76 4.36 ± 0.32 < 0.1 <0.1 pU702 SKK896 <0.1 <0.1 <0.1 <0.1 pKKl111 SKK821 52.86 ±4.16 5.76 ± 0.66 13.76 ± 1.74 0.65 ± 0.16 pKK1111 SKK896 16.05 ± 0 93 5.23 ± 0.67 7.28 ± 1.66 1.38 ± 0.20

+ Histidine, CAA medium containing 10 mM L-histidine; - Histidine, glutamate minimal medium. 108 High-resolution S1 nuclease analysis showed that although the hutH transcription initiation site in SKK896 is identical to that in the wild-type strain (Fig. 24), the level of transcript in the mutant strain is reduced approximately 10-fold. The sequence of 369 nt extending from 291 nt upstream to 78 nt downstream of the beginning of the hutH coding sequence in SKK896 is also identical to that of SKK821 (data not shown). These results indicate that there is less expression of hufH in SKK896, but that this is not the consequence of an alteration in the upstream regulatory region or the promoter.

Repressor titration experiments showed that when grown in the absence of L-histidine,

SKK896 containing multiple copies of wild-type DNA extending 333 nt upstream of the hufH coding sequence produced a significant level of urocanase (Table 5). This result suggested that the negative regulatory factor in SKK896 is functional and multiple copies of wild-type upstream DNA can effectively sequester the negative regulatory factor in

SKK896. CHAPTER IV

DISCUSSION

A 3.45 kb EcoRI/W/ndlll fragment from S. griseus was cloned by plaque hybridization with an oligonucleotide probe derived from the amino acid sequence of the

N-terminus of purified histidase, followed by subcloning. This fragment contains two streptomycete open reading frames (ORFs; Fig. 15). Identification of ORF2 as the histidase structural gene was made on the basis of: (1) the deduced amino acid sequence identity of the N-terminus of the ORF2 gene product with the N-terminus of the purified protein (Fig. 14; Wu et al. 1992); (2) similarity of the deduced molecular mass of the ORF2 gene product with that of the purified protein (Wu et al., 1992); (3) similarity of the deduced amino acid sequence of the ORF2 gene product with those of other bacterial and mammalian histidases (Fig. 17); and (4) complementation of a Hut' mutant deficient in histidase activity (Table 4).

The deduced amino acid sequence of histidase from S. griseus shows homology to those of histidase from other bacteria and mammals as well as plant and fungal phenylalanine ammonia-lyase. A phylogenetic analysis of the evolutionary relatedness of amino acid sequences of histidases revealed that 5. griseus histidase is less related to other bacterid and mammalian histidases. Surprisingly, rat histidase is more closely related to histidase from B. subtilis and P. putida than is histidase from S. griseus.

109 110

Perhaps the divergence of streptomycete histidase is a reflection of the distinct characteristics of the enzyme, including hysteresis (Kroening and Kendrick, 1987), the

absence of a requirement for divalent cations or thiol reagents for maximal activity

(Kroening and Kendrick, 1987; Wu et al., 1992), the higher affinity of histidase for its substrate (Wu et at., 1992; White and Kendrick, 1993), and the potent competitive inhibition by histidinol phosphate (Wu et al., 1992; White and Kendrick, 1993).

DehydroaJanine (Consevage and Phillips, 1985), cysteine (Klee and Gladner,

1972), and arginine residues (White and Kendrick, 1993) have been implicated in the catalytic properties of histidase. Inspection of the amino acid sequences of four histidases (Fig. 17) revealed that there are 11 conserved arginines and one conserved cysteine residue. Interestingly, the sole conserved cysteine residue (Cys^, in 5. griseus histidase) is adjacent to a conserved arginine residue (Arg^,). The serine residue identified by Hernandez et al. (1993) is at position 146 in S. griseus histidase (Fig. 17).

There are 10 additional conserved serine residues in the four histidases (Fig. 17), any one of which may be the source of dehydroalanine.

The guanine and cytosine (G+C) content of the histidase structural gene is 74%, characteristic of the average G+C content of streptomycete DNA (Chater and Hopwood,

1984). Despite this overall high G+C content in hutH , the G+C content at the N-terminal coding region of the hutH gene is relatively low. I was fortunate that all possible codons, not just those containing G or C in the third codon positions, were included in the degenerate oligonucleotide probe. Otherwise, a biased oligonucleotide probe would likely have failed to detect the hutH gene from the S. griseus genomic DNA library. 111 An open reading frame (ORF1) that would be transcribed in the same direction as hutH lies upstream of hutH. Although the 5' end of 0RF1 is deleted in pKK1100, ORF1 is likely to be complete in pKK1111. The amino acid sequence deduced from the available nucleotide sequence of ORF1, which would generate a polypeptide of Mr >

25,400, resembles the sequence of the putative product of ORF183 from 5. ambofaciens

(Hagfcge et al., 1993, and unpublished). As a gene that maps to an integrating plasmid, pSAM2, ORF183 is thought to be involved in the regulation of pSAM2 replication. Recent circumstantial evidence (K.V. Srinivasan, unpublished) suggests that the gene product of ORF1 may be involved in histidine utilization.

There is no evidence that the 0.5 kb DNA downstream of hutH contains a streptomycete open reading frame or a tRNA structural gene. In other bacterial systems

(Chasin and Magasanik, 1968; Hu and Phillips, 1988; Magasanik, 1978), the hutH gene is part of an operon and is co-transcribed with hutU. The results of low-resolution S1 mapping clearly showed that the hutH transcript in S. griseus is monocistronic (Fig. 23).

Thus, the hutH gene is not part of an operon, and the hut genes of 5. griseus are organized differently from those of other bacteria

S1 mapping analysis showed that the hutH transcript is 1.6 kb in length (Fig. 23) and initiates at the adenine that marks the ATG translation initiation codon (nt 835; Fig.

24). Three nucleotides downstream of the hutH coding sequence is an inverted repeat with a AG of -42.9 Kcal/mol. It appears that the hutH transcript terminates or is processed to a 3' endpoint in the neighborhood of this inverted repeat. Further downstream of hutH is A-like DNA (nt 2947 through 3444), whose nucleotide sequence 112

is identical to phage X DNA within the int gene (Hendrix et al., 1983). Although Southern

analysis of S. griseus genomic DNA using phage X DNA (containing the int gene) as

probe suggested that S. griseus does not contain A-like DNA (data not shown), other

experiments (Fig. 20) suggested that there were A-like sequences in S. griseus genomic

DNA. For two reasons, however, I hypothesize that the A-like DNA in the hutH clones

originated from AEMBL4 and is therefore a cloning artefact. First, AEMBL4 contains the int and xis genes (Frischauf et al., 1983) and was used for the construction of the S. griseus genomic DNA library. Second, computer searches of the GenBank database

revealed that several eucaryotic gene sequences, such as one describing the sequence

of a protein from Plasmodium falciparum (Lenstra et al., 1987) and one describing a

chicken cardiac phospholamban gene (Toyofuku and Zak, 1991), contain sequences that are identical to the A-like DNA in the hutH clone. In these two studies, either AEMBL4 or

AEMBL3 was used for the construction of the DNA libraries.

If my hypothesis is correct, then some level of DNA rearrangement occurred during the cloning of the hutH gene. Despite the apparent insertion of X DNA into the original cloned fragment, the region extending from nt 1 to nt 2938 (the rightmost Sa/I site in pKK540) showed no evidence of gene rearrangement (Figs. 20 and 21). PCR amplification of S. griseus genomic DNA should confirm the absence of A-like DNA downstream of hutH. Cloning and nucleotide sequence determination of DNA downstream of the hutH gene in S. griseus wilt be useful to answer: (1) why did recombination of phage X DNA and S. griseus DNA occur?; (2) what is the nucleotide sequence of chromosomal DNA at the region in which X DNA was found in the hutH clones?; and (3) are there any other hut genes downstream of hutH? 113 Southern analysis of S. griseus genomic DNA using the hutH clone as probe suggested that S. griseus possesses only one copy of hutH that is similar to the one I have cloned (Fig. 21). A parallel experiment using the same probe hybridized with S. coelicoior chromosomal DNA suggested that the hutH-like sequence is present in S. coeiicolor (Fig. 21), in agreement with enzymological and genetic data (Kendrick, 1979;

Kendrick and Wheelis, 1983).

Regions at 10 nt and 35 nt preceding the hutH translation initiation codon resemble a streptomycete consensus promoter which would be expected to be active during vegetative growth (Fig. 25; Strohl, 1992). Promoter-probe studies using plJ486 and plJ487 as well as pXE4 showed that a DNA fragment extending 67 bp upstream of the hufH translation start codon has promoter activity (Fig. 22, Tables 6 and 7). To classify streptomycete promoters, Strohl analyzed 119 streptomycete promoter sequences (Strohl, 1992). Of the promoters investigated, 28 were classified into a group whose sequences resemble the consensus sequence for E. coli promoters recognized by RNA polymerase containing the a70 subunit (Ea70). On the basis of nucleotide sequence similarity, the hutH promoter belongs to this group and is likely to be recognized by an Ec^-type of RNA polymerase, the major class of RNA polymerase found in S. coelicoior (Buttner et al., 1990; Tanaka et al., 1988). Buttner et al. (1990) have isolated null mutants of S. coelicoior that are defective in three of the four hrd genes, which encode aTO-like sigma factors. The introduction of the hutH: x/!E transcriptional fusion into these mutant strains should lead to the identification of the hrd gene that is required for transcription of hutH. Fig. 25. Consensus sequences of E co/Mike streptomycete promoters and the

nucleotide sequence upstream of hutH from S. griseus. The nucleotide

sequences of 28 streptomycete promoters resemble the consensus

sequence for E. coli promoters recognized by E an (Strohl, 1992). Capital

letters denote the most conserved nucleotides; the less conserved

nucleotides are denoted by lower case letters. Transcription initiation sites

are designated *+1*. The minor transcription initiation site of hutH is

marked with a small m+V.

114 115

-35 -10 +1 Consensus TTGaCa.,17nt. .TATAAT. .5-6nt. .C

-35 -10 +1 Streptomyces TTGACg.. 17nt. .TAgaaT. ,6-7nt. .G Consensus a egg c

. _ -35 -10 +1 (Sgriseus) GTGACA. .17nt. .TACGCT. .7nt. . ATGGATATG

Figure 25 116 The promoter-probe vectors plJ486 and plJ487 contain a promoterless neo gene that confers resistance to kanamycin (Ward et ai., 1986). There are two drawbacks to measuring S. griseus hutH promoter strength using these two vectors. First, Babcock and Kendrick (1990) observed that neither plJ486 nor plJ487 can be maintained as a high copy number vector in S. griseus. By visual comparisons with the low copy number vector pXE4 from S. griseus (1-2 copies per chromosome; K. E. Kendrick, unpublished data) on an agarose gel, I estimated the copy number of these two vectors in S. griseus to be less than one. The vectors plJ486 and plJ487 contain both a replicon and a region for plasmid stability from plJ101 found in S. IMdans ISP 5434 (Kieser et al., 1982).

Although the stability of plJ101 in S. griseus has not been tested, a derivative of plJ101

(plJ303) that retains both the replicon and stability regions of plJI 01 was structurally stable in S. IMdans 66 but not in S. griseus ATCC 10137 (Kieser et al., 1982). In contrast, the streptomycete vector plJ702, also containing the replicon and stability region of plJ101 (Katz et al., 1983), is stable and maintained at high copy number in S. griseus (50 per chromosome; K.E. Kendrick, unpublished data). Thus, there may be a factor other than the replicon that causes instability of plJ486 and plJ487 in S. griseus. Second, the assessment of S. griseus promoter activity in a heterologous, high copy number system

(S. IMdans TK24) may not reflect the real promoter strength in S. griseus. The copy number of plJ486 and plJ487 in S. IMdans TK24 has been reported to be between 100 and 200 per chromosome (Ward et al., 1986). Thus, differences in the levels of resistance between S. griseus and S. IMdans transformants are likely to be due to the copy number of these vectors in each species (Table 6). An enzyme-linked immunosorbent assay for the quantitation of neomycin phosphotransferase II, the product 117 of neo, may provide a more sensitive and accurate measure of promoter activity than the more subjective assessment of resistance to kanamycin.

Because instability of plJ486 and plJ487 was inherent in S. griseus, another promoter-probe vector, pXE4, was used. The plasmid pXE4 was constructed by Ingram et al. (1989) and has been used successfully in their study of regulation of galPl expression in S. coelicoior as well as the regulated expression of chi63, a gene that encodes chitinase in Streptomyces plicatus (Delic et al., 1992). This vector contains a promoterless xyiE gene originally isolated from P. putida plasmid pWWO (Worsey and

Williams, 1975). The product of xylE, catechol 2,3-dioxygenase, catalyzes the oxidation of colorless catechol to a yellow product, 2-hydroxymuconic acid semialdehyde (Bayly et al., 1966). pXE4 is stable in S. griseus and is a low copy number vector in

Streptomyces (Ingram et al., 1989). The copy number of plJ922 (which uses the same replicon as pXE4; Lydiate et al., 1985) in 5. griseus is 1-2 per chromosome (K. E.

Kendrick, unpublished data), characteristic of plasmids containing the SCP2 replicon.

Promoter activity can be quantified by measuring XylE activity in a simple spectrophotometric assay. In addition, pXE4 is a bifunctional plasmid with a ColE1 origin of replication to facilitate plasmid construction and screening in E. coli cells rather than in more slowly growing Streptomyces cells.

The results of promoter-probe studies using pXE4 also indicated that the hutH promoter lies within 67 bp upstream of the hutH coding sequence (Table 7). Low background levels of XylE activity were observed, consistent with those found in 5.

IMdans containing pXE4 (Ingram et al., 1969). The levels of XylE activity may reflect 118 accurately the S. griseus hutH promoter strength, but, unlike the results obtained using plJ486 and plJ487, there is no evidence for enhanced expression of hutH in the presence of upstream sequence. By using different plasmids (i.e., not pXE4) constructed with xylE as a reporter gene for promoter-probe studies of dagA and amyL, M. J. Bibb observed that the levels of promoter activity were low, and he was unable to detect glucose repression of either gene despite his earlier observations that these genes contain relatively strong promoters and are subject to glucose repression (personal communication). A recent report concerning the inactivation of XylE indicated that after oxidation of catechol by

XylE, XylE is reduced and inactivated (Polissi and Harayama, 1993). The product olxyFT, which maps immediately upstream of and is co-transcribed with xylE in P. putida plasmid pWWO (Harayama and Rekik, 1990), is responsible for oxidation (and consequent activation) of the inactive XylE (Polissi and Harayama, 1993). Although the xyfT gene is absent from Bibb’s xylE plasmid constructions (personal communication), examination of the genetic map and origin of pXE4 revealed that xyU is present in pXE4 (Appendix

A; Harayama and Rekik, 1990; Ingram et al., 1989; Zukowski et al., 1983). Therefore, it is unlikely that the accumulation of inactive XylE explains the low apparent level of hutH promoter activity in these studies.

There are several genes that have been used as reporter genes for the construction of Streptomyces promoter-probe vectors. These gene products have features of pigment production, light emission, antibiotic resistance, or enzyme activity when a promoter-active fragment is inserted in the orientation that directs transcription of the reporter gene. These reporter genes include the actinorhodin biosynthetic genes from 5. coelicoior that confer production of a brown pigment (Horinouchi and Beppu, 119

1985), lux, which encodes luciferase from Vibrio harveyi (Sohaskey et al., 1992), neo,

which confers kanamycin resistance from Tn5 (Daza et al., 1991; Ward et al., 1986), mdh,

which encodes the extremely thermostable malate dehydrogenase from Thermus flaw s

(Vujaklija et al., 1991), and xylE, which encodes catechol dioxygenase from P. putida

plasmid pWWO (Ingram et al., 1989; Kuhstoss and Rao, 1991). Of these promoter-probe

vectors, assessment of promoter strength by production of pigment is not a quantitative

measure and can be complicated by endogenous pigments produced by the host strain.

The level of expression of lux in the promoter-probe vectors that rely on this reporter gene

is low; site-directed mutagenesis is being undertaken to improve the expression (KF.

Chater, personal communication). A similar problem occurs in using the mdh gene: the level of malate dehydrogenase activity was only two-fold greater than background when the promoter-active fragment was in the correct orientation relative to the reporter gene

(Vujaklija et al., 1991). Thus, to date it appears that there is not a universally suitable

Streptomyces promoter-probe vector with a sensitive and simple detection system as well as a faithful reflection of promoter strength and regulation.

Determination of the N-terminal amino acid sequence of histidase purified from

S. griseus by Edman degradation analysis revealed that the hutH translation initiation site corresponds to nt 835 (Figs. 5 and 14). Promoter-probe studies and the presence of a promoter consensus sequence suggested that the hutH promoter may lie Immediately upstream of the hutH translation initiation site. There is no obvious ribosome binding site

(Bibb and Cohen, 1982; Hopwood et al., 1986) upstream of the translation initiation site, suggesting the existence of coincident initiation of transcription and translation, translational coupling, or a weak ribosome binding site. High-resolution S1 mapping 120 confirmed that the hutH gene in $. griseus is expressed as a leaderless transcript (Fig.

24). Studies of translation of leaderless messages of the aminoglycoside phosphotransferase gene (aph) from S. fradiae indicated that streptomycete ribosomes can efficiently recognize and initiate translation at the 5' terminal AUG codon of a transcript in the absence of a conventional Shine-Dalgamo interaction between 16S rRNA and mRNA (Janssen et al., 1989; Jones et al., 1992).

A minor transcript initiating 6 nucleotides downstream of the major site starts at the adenine of the AUG that encodes the third amino acid of purified histidase (Fig. 24).

According to the survey of 28 streptomycete E. co/Mike promoters (Strohl, 1992), the most common distance between the -10 hexamer and the transcription start site is 7 nucleotides, which is in agreement with the transcription start site of the major hutH transcript. It is possible that the minor transcript is a processed product of the major one. The N-terminal amino acid sequence analysis did not, however, show detectable amounts of the shorter polypeptide. The insertion of 4 nucleotides (ATGC) immediately upstream of the transcription start site (ATG) of the aph gene to generate a potential frameshift (ATGCATG) resulted in transcription initiating only at the upstream start site in vivo (Jones et al., 1992). Capping analysis (Shuman and Hurwitz, 1982) should clarify whether or not the minor transcript is the result of a processing event or signifies a true transcription initiation site.

To date, at least 12 genes in actinomycetes are known to have coincident initiation of transcription and translation (Ishizuka et al., 1992; Strohl, 1992). Of these 12 genes,

10 are involved in secondary metabolism such as antibiotic production or antibiotic resistance. For example, afsA, an A-factor biosynthetic gene from S. griseus, is required for the synthesis of A-factor which is involved in streptomycin production (Horinouchi et al., 1989), and aph from S. fradiaa confers resistance to neomycin (Janssen et al.,1969).

The other two genes include korB, which is involved in regulation of transfer of the

Streptomyces plasmid plJ101 (Stein et al., 1989), and cdh, which encodes cholesterol dehydrogenase in Nocardia species (Horinouchi et al., 1991). Discovery of the unusual gene structure of hutH from S. griseus confirms that coincident initiation of transcription and translation is not unique to genes involved in secondary metabolism but is also present in streptomycete genes required for primary metabolism. Other procaryotic genes having this structure include the X c l gene transcribed from the pRM promoter

(Ptashne et al., 1976), the tetR gene in 7n1721 (Klock and Hillen, 1986), and the bacteriorhodopsin (bop; Dunn et al., 1981), bacteriorhodopsin-related (brp; Betlach et al.,

1984), and halo-opsin (hop; Blanck and Oesterheit, 1987) genes in Haiobacterium halobium.

Low-resolution S1 nuclease analysis indicated that expression of hutH is regulated at the level of transcription, and synthesis of hutH mRNA is inducible (Fig. 23). Under non-inducing growth conditions, SKK821 transformants containing multiple copies of a

DNA fragment extending 176 nt but not a DNA fragment extending 67 nt upstream of the hutH transcription start site showed relatively high levels of histidase and urocanase activities (Table 5). These results suggested that the 176 bp fragment present at high copy number is able to titrate a diffusible negative regulatory factor. Computer analysis of the 176 bp region revealed an inverted repeat (nt 750 through nt 793) with a AG of

•23.7 Kcal/mol. The 176 bp but not the 67 bp upstream region used for repressor 122 titration experiments contains this complete inverted repeat. I favor the hypothesis that the inverted repeat centered at -63 relative to the transcription start site of hutH functions as a repressor binding site. The nucleotide sequence of this inverted repeat is not similar to those of the repressor binding sites of the hut genes that are centered at the -35 region in K. aarogenes (Schwacha and Bender, 1990) and P. putida (Hu et al., 1989). Because there is evidence of co-expression of hutH and hutU in S. griseus in response to the same negative regulatory factor (Tables 4 and 5), comparison of the nucleotide sequence upstream of hutU to that upstream of hutH would be likely to identify the binding site for the repressor that appears to co-regulate hutH and hutU. To test whether this inverted repeat is involved in negative control of hut expression, PCR-generated fragments containing the entire inverted repeat or portions of this repeat could be examined for their ability to elicit repressor titration.

As in the enteric bacteria (Schiesinger et al., 1965) and pseudomonads (Hu et al.,

1989), urocanate but not L-histidine is the physiological inducer for hutH expression in

S. griseus (Kroening and Kendrick, 1989). With respect to the fate of urocanate, histidase is an inducer-producing enzyme, and urocanase is an inducer-destroying enzyme

(Magasanik, 1978). It is therefore important for an organism to co-regulate the activities of these two enzymes to guarantee that L-histidine can be utilized efficiently. This goal can be achieved by coordinate expression of hutH and hutU in two ways. In the case of the enteric bacteria (Magasanik, 1978) and pseudomonads (Hu and Phillips, 1988), hutH and hutU are arranged in an operon. In the case of S. griseus, hutH and hutU are not co­ transcribed but are apparently subject to the same negative regulatory mechanism. This would ensure that expression of hutH is accompanied by expression of hutU. 123

There is physiological significance to the low level of hutH expression in the absence of the inducer. Because urocanate is the inducer, a cell must be able to convert exogenous L-histidine to urocanate in order to induce the hut genes. Studying histidine utilization in P. putida, Hu et al. (1969) have noted that a low level of histidase is always present. They hypothesized that in the presence of L-histidine, urocanate accumulates from conversion of L-histidine by the low level of existing histidase and rises to a level sufficient for induction. In P. putida, hutH is part of the hutUHIG operon, and the promoter of this operon immediately precedes hutU (Hu and Phillips, 1988). The nucleotide sequence of hutH from P. putida revealed a potentially weak promoter immediately upstream of hutH (Consevage and Phillips, 1990), but whether this region functions as a bufW-specific promoter is still unknown. It is likely that the same hypothesis applies to hut induction in S. griseus. The results of low-resolution S1 mapping provide the first evidence to support the hypothesis of Hu et al. (1989) and are consistent with enzymological data that demonstrate low histidase activity in the wild-type strain grown in a non-inducing medium (Kroening and Kendrick, 1989).

By analysis of 76 E. coii promoter regions, Collado-Vides et al. (1991) established a general rule that describes the degree of repression and location of the repressor binding site relative to the promoter. These repressor binding sites include: (1) upstream of the -35 region, (2) the spacer region between -35 and -10, and (3) the -10 region overlapping the transcription initiation site. Of these binding sites, a repressor bound to the spacer region is considered to be the most effective in preventing RNA polymerase binding because the -35 and -10 regions are completely blocked by the repressor

(Collado-Vides et al., 1991). A repressor bound at the -10 region occludes the critical 124 transcription initiation site but leaves the -35 region potentially available for RNA polymerase to form a productive RNA-DNA complex. On the other hand, when the repressor binding site lies upstream of the -35 region, the critical -10 region and transcription initiation site are largely available for access of RNA polymerase to this region; this situation is believed to confer the weakest repression (Collado-Vides et al.,

1991) and would be predicted to result in a low level of gene expression under non­ inducing conditions. Because there is a low level of expression of hutH in 5. griseus grown in the absence of inducer, it is reasonable to speculate that the repressor binding site of hutH in S. griseus is located upstream of the hutH promoter as proposed.

The DNA fragments used in the promoter-probe studies and repressor titration experiments also include approximately 85 nucleotides of the N-terminal coding region of hutH (Fig. 22). Analysis of the nucleotide sequence surrounding the hutH transcription/translation start site revealed that in addition to the proposed repressor binding site, another inverted repeat with a AG of -20.7 Kcal/mol lies 15 nucleotides downstream of the transcription/translation initiation site (Fig. 14). Promoter-probe studies using plJ486 and plJ487, but not pXE4, showed that the promoter activity increased with increasing length of the hutH upstream DNA (from 176 bp to 291 or 333 bp; Tables 6 and 7). This observation was most evident in the S. IMdans transformants.

These results suggest that there may be an additional regulatory region upstream of the hutH coding sequence. It is possible that the inverted repeat immediately upstream of the transcriptional fusion junction in pXE4 as well as plJ486 and plJ487 may reduce expression of the reporter genes by acting as a weak transcription terminator. Deletion 125 or mutation analysis would resolve whether this downstream inverted repeat can affect hutH expression in vivo.

A Hut mutant of S. griseus (SKK896) unable to utilize L-histidine as sole source of nitrogen was originally isolated by T. A. Kroening in our laboratory. SKK896 showed no histidase and urocanase activities when grown in the presence of L-histidine, but the urocanase activity was detected when the mutant strain was grown in the presence of urocanate (Kroening and Kendrick, 1989). These results suggested that SKK896 is defective in the production of active histidase. A 4.0 kb BamH\ fragment containing the hutH allele and 1.5 kb of adjacent upstream DNA from SKK896 was cloned. SKK896 harboring multiple copies of this flamHI fragment in (rans produced substantial levels of both histidase and urocanase when grown in the presence of L-histidine (Table 8), suggesting that SKKB96 is defective either in the hutH coding sequence to become a less active gene product or in the regulation of hutH expression or enzymatic activity.

Because enzymological studies showed that SKK896 does not produce any detectable histidase (Kroening and Kendrick, 1989), the former possibility is less likely.

Complementation studies showed that SKK896 containing multiple copies of the wild-type allele of hutH produced high constitutive levels of histidase (Table 4). These results suggest that SKKB96 is unlikely to have a mutation in the histidase structural gene. It is also unlikely that SKK896 is defective in the protein that activates histidase: whereas the wild-type histidase undergoes spontaneous activation during the ten-minute course of a histidase assay, there is no detectable histidase activity in extracts prepared from SKK896 and assayed under the same conditions. 126 If the mutation in SKK896 does affect the regulation of hutH expression, one would

expect that the level of hutH transcription or translation would be reduced. The S1

mapping studies showed that this was the case for SKK896. One mechanism by which

transcription could be reduced is by mutation in the upstream regulatory region, which

would be expected to reduce binding of a positive regulatory factor or RNA polymerase,

or, alternatively, to enhance binding of a negative regulatory factor. The nucleotide

sequence of the DNA upstream of hutH in SKK896 is identical to the wild-type sequence.

This region includes the proposed negative regulatory factor binding site. It seems, therefore, that the defect in SKK896 more likely resides in a regulatory factor. A summary

of the evidence leading to this conclusion is presented in Fig. 26. Consistent with this

interpretation, recent results (K.V. Srinivasan, unpublished) suggest that SKK896 contains a mutation within a gene that maps a short distance upstream of hutH. It is tempting to speculate that ORF1, which resembles a reported regulatory protein, may be involved in histidine utilization and be the site of the mutation in SKK896. Future experiments will be designed to test this possibility. Figure 26. Evidence indicating the nature of the mutation in SKK896.

127 128 SKK896 (absence of histidase activity - Table 4)

possible mutations

Mutation in Mutation In M utation in the regulation of regulation of histidase histidase activity hutH expression structural gene

^ unlikely l ik e l y ^ unlikely

Evidence: absence of Evidence: abnormal Evidence: suppression spontaneous activation regulation of hi/fH in of hutH In SKK896 of histidase from SKK896 transformed with transformed with the SKK896 during reaction the wild-typehutH hutH gene from SKK896 (Table 8)

In regulatory site In gene encoding (i.e., cfc-acting) a regulatory factor (I.e., frans-acting)

^ unlikely ^ likely

Evidence: (1) identity Evidence: reduced of DNA sequence levels of hutH mRNA extending 290 nt upstream of hutH In SKK896 with that in the wild-type strain; (2) normal repressor ______titration characteristics _J Conclusion: SKK896 is most likely defective in a factor involved in the regulation of hutH expression

Figure 26 129

Summary

Comparisons of the histidase primary structures and organization of hutH among organisms suggest that hutH from S. griseus may have arisen via a divergent evolutionary pathway. Unlike hutH gene expression in Pseudomonas, the enteric bacteria, and

Bacillus, the transcript of the hutH gene in S. griseus is monocistronic and leaderless.

Like hutH expression in other bacteria, expression of the hutH gene in S. griseus is under negative control. Positive regulation may also play a role in hutH expression. Analysis of a Hut' mutant suggests that it is defective in a factor that regulates the expression of hutH. LIST OF REFERENCES

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Map# of Plasmid Vector* Used in This Study

140 Fig. 27. Circular map of pGEM-4Z. This E. coli transcription vector contains SP6

and T7 promoters as well as the a peptide of /7-galactosidase (lacZ) and

the ampicillin resistance gene (amp).

141 pGEM-4z BamH\ 2746 bp

FcoRI

Figure 27 Fig. 28. Circular maps of pUC18 and plJ2925. These two E. coli vectors contain

the a peptide of /Ngalactosidase (/acZ) and the ampicillin resistance gene

(amp). The vector plJ2925, derived from p(JC18, contains additional

restriction sites in the polylinker region.

143 144 SalG\ B gl\ fc o R I Ssfl S m d BamHI Xbck HhcW W/odlll PsH BgH\ PIJ2925 HhdM Pstl HncM X b d fidmHI S n d Ssfl pUC18 I I

pUC18 or PIJ2925

a m p

Figure 28 Fig. 29. Circular map of plJ702. This Streptomyces vector contains the tyrosinase

genes (melCI and me/C2) for white/brown color detection and the

thiostrepton resistance gene (tsr) for antibiotic selection. This high copy

number vector is a derivative of piJ101 (Hopwood et al., 1985; Katz et al.,

1983).

145 146

Ps1\

m e lC l Bgli\ PIJ702 5.8 kb

m e\C2

tsr

Figure 29 Fig. 30. Circular map of plJ486 and plJ487. The vector plJ486 contains a

promoterless neo gene from Tn5 that confers kanamycin resistance for

promoter-probe studies. This vector contains the bacteriophage fd

transcription terminator (stem-and-loop structure) upstream of the multiple

cloning site and the thiostrepton resistance gene (tsr) for selection. The

vector plJ487 is identical to plJ486 except that the restriction sites in the

poly linker region are in opposite orientation relative to the neo gene (Ward

et al., 1986).

147 148

PIJ486

H/nd

SomHI PIJ486/487 6.2 kb fcoRI

Figure 30 Fig. 31. Circular map of pXE4. This vector contains a promoterless xylE gene

encoding catechol 2,3-dioxygenase from Pseudomonas putida. This

vector is a bifunctional plasmid containing the ColE1 origin for replication

in E. coli and the SCP2 replicon to maintain 1 -2 copies per chromosome

in Streptomyces. Also present are the thiostrepton gene (tsr) for selection

in Streptomyces and the ampicillin resistance gene (amp) for selection in

E. coli. The bottom portion of the figure depicts the physical map of the

xylE and xytT genes in the vector (Ingram et al., 1989).

149 150

BarriHl

xylTE

SCP2 rep

pXE4 colEI 17.5 kb rep

a m p

tsr

BamH\ xyU xylE

4 5 0 b p 9 2 0 b p ------350 b p *

Figure 31 Appendix B

List of Plasmids Constructed In This Study

151 Table 9. List of plasmids constructed.

The 8 kb EcoRI/Bglll fragment from (lambda)KK500 ligated into pKK500 PGEM-4Z EcoRI-BamHI-digested pGEM-4Z 8 kb 544-3444 E. coli TB1 The 2.5 kb EcoRI/BamHI fragment from (lambda|KK501 ligated into PKK501 PGEM-4Z EcoRI-BamHI-digested pGEM-4Z 2.48 kb 1-2476 E. coli TB1

pKK502 PGEM-4Z Identical to pKK501 2.48 kb 1-2476 E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-2267 PKK503 PGEM-4Z III, blunt-ended with SI nuclease, and religated 2.3 kb lapprox.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK504 PGEM-4Z III, blunt-ended with SI nuclease, and religated 2.2 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-2039 pKK505 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 2.0 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK506 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 0.4 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-1500 pKKS07 pGEM-4Z III, blunt-ended with S1 nuclease, and religated 1.5 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-1201 pKK508 pGEM-4Z III, blunt-ended with S1 nuclease, and religated 1.2 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-1117 pKK509 pGEM-4Z III, blunt-ended with S1 nuclease, and religated 1.12 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK510 pGEM-4Z III, blunt-ended with SI nuclease, and religated 0.1 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-1737 pKK511 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 1.74 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK512 pGEM-4Z III, blunt-ended with SI nuclease, and religated 1.5 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease PKK513 DGEM-4Z III. blunt-ended with S1 nuclease, and religated 0.5 kb n.d. E. coli TB1 Table 9, cont.

pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-669 pKK514 pGEM-4Z III, blunt-ended with SI nuclease, and religated 0.67 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK515 PGEM-4Z III, blunt-ended with SI nuclease, and religated 1.05 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK516 pGEM-4Z III, blunt-ended with SI nuclease, and religated 1.7 kb n.d. E. coli TB1 pKKSOI was digested with Xbal and Pstl, digested with Exonuclease 1-1867 pKK517 PGEM-4Z III, blunt-ended with SI nuclease, and religated 1.87 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK518 PGEM-4Z III, blunt-ended with S I nuclease, and religated 1.8 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK519 PGEM-4Z III, blunt-ended with S I nuclease, and religated 0.9 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-840 pKK520 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 0.84 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK521 pGEM-4Z III, blunt-ended with S I nuclease, and religated 0.8 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-1410 pKK522 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 1.41 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK523 pGEM-4Z III, blunt-ended with SI nuclease, and religated 1.2 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK524 PGEM-4Z III, blunt-ended with SI nuclease, and religated 0.9 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease 1-2361 pKK525 pGEM-4Z III, blunt-ended with SI nuclease, and religated 2.36 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease PKK526 PGEM-4Z III, blunt-ended with SI nuclease, and religated 2.3 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease PKK527 PGEM-4Z III, blunt-ended with SI nuclease, and religated 2.3 kb n.d. E. coli TB1 Table 9, cont.

pKK501 was digested with Xbal and Pstl, digested with Exonuclease PKK528 pGEM-4Z III, blunt-ended with S1 nuclease, and religated 2.3 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK529 pGEM-4Z III, blunt-ended with SI nuclease, and religated 0.5 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease PKK530 PGEM-4Z III, blunt-ended with S i nuclease, and religated 0.75 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonudease PKK531 pGEM-4Z III, blunt-ended with S I nuclease, and religated 0.3 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonudease 1-364 PKK532 pGEM-4Z III, blunt-ended with S I nuclease, and religated 0.37 kb (approx.) E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonudease pKK533 PGEM-4Z III, blunt-ended with SI nuclease, and religated 0.25 kb n.d. E. coli TB1 pKK501 was digested with Mlul and Hindlll to excise the 0.67 kb Mlul/Hindlll fragment and replaced with the 1.64 kb Mlul/Hindllt PKK534 PGEM-4Z fragment from pKK500 3.45 kb n.d. E. coli TB1 pKK501 was digested with Mlul and Hindlll to excise the 0.67 kb Mlul/Hindlll fragment and replaced with the 1.64 kb Mlul/Hindlll PKK535 pGEM-4Z fragment from pKK500 3.45 kb n.d. E. coli TB1 pKK501 was digested with Mlul and Hindlll to excise the 0.67 kb Mlul/Hindlll fragment and replaced with the 1.64 kb Mlul/Hindlll E. coli PKK536 PGEM-4Z fragment from pKK500 3.45 kb n.d. JM109 pKK501 was digested with Mlul and Hindlll to excise the 0.67 kb Mlul/Hindlll fragment and replaced with the 1.64 kb Mlul/Hindlll E. coli PKK537 PGEM-4Z fragment from pKKSOO 3.45 kb n.d. JM109 Table 9, cont.

The 3.45 kb EcoRI/Hindlll fragment from pKK535 was blunt-ended with S I nuclease and inserted into the Smal site of pUC18. The direction of transcription of hutH in this construction is from toward pKK538 pUC18 the Hindlll site in the poiylinker of the vector. 3.45 kb 1-3444 E. coli TB1

The 3.45 kb EcoRI/Hindlll fragment from pKK535 was blunt-ended with S I nuclease and inserted into the Smal site of pUC18. The direction of transcription of hutH in this construction is from toward pKK539 PUC18 the Hindlll site in the poiylinker of the vector. 3.45 kb 1-3444 E. coli TB1 The same insert as described in pKK538 was ligated into the Smal pKK540 PUC18 site of the vector in the opposite orientation. 3.45 kb 1-3444 E. coli TB1 The same insert as described in pKK538 was ligated into the Smal PKK541 pUC18 site of the vector in the opposite orientation. 3.45 kb 1-3444 E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease PKK542 pGEM-42 III, blunt-ended with S1 nuclease, and religated 1.6 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK543 pGEM-4Z III, blunt-ended with S I nuclease, and religated 0.8 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease PKK544 PGEM-4Z III, blunt-ended with S I nuclease, and religated 0.1 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK545 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 1.65 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK546 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 0.85 kb n.d. E. coli TB1 pKK501 was digested with Xbal and Pstl, digested with Exonuclease pKK547 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 1.1 kb n.d. E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2839 PKK548 PUC18 III, blunt-ended with S1 nuclease, and religated 2.84 kb (approx.) E. coli TB1 Table 9, cont.

pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2595 pKK549 pUC18 III, blunt-ended with S1 nuclease, and religated 2.6 kb (approx.) E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease PKK550 PUC18 III, blunt-ended with SI nuclease, and religated 2.65 kb n.d. E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2717 pKK551 pUC18 III, blunt-ended with S1 nuclease, and religated 2.72 kb (approx.) E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2568 pKKS52 pUC18 III, blunt-ended with S1 nuclease, and religated 2.57 kb (approx.) E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2408 PKKS53 PUC18 III, blunt-ended with S1 nuclease, and religated 2.41 kb (approx.) E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 1368 (approx.) pKK554 PUC18 III, blunt-ended with S1 nuclease, and religated 2.08 kb -3 4 4 4 E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2202 pKK555 pUC18 III, blunt-ended with SI nuclease, and religated 2.2 kb (approx.) E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease PKK556 PUC18 III, blunt-ended with SI nuclease, and religated 2.3 kb n.d. E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease PKK557 pUC18 III, blunt-ended with S1 nuclease, and religated 2.55 kb n.d. E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2500 PKK558 PUC18 III, blunt-ended with SI nuclease, and religated 2.5 kb (approx.) E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 1580 (approx.) pKK559 PUC18 III, blunt-ended with S1 nuclease, and religated 1.87 kb -3 4 4 4 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 1752 (approx.) PKK560 PUC18 III, blunt-ended with S1 nuclease, and religated 1.69 kb -3 4 4 4 E. coli TB1 Table 9, cont.

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 1889 (approx.) pKK561 pUC18 III, blunt-ended with S1 nuclease, and religated 1.56 kb •3 4 4 4 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 2138 (approx.) pKK562 pUC18 III, blunt-ended with S1 nuclease, and religated 1.31 kb - 3444 E. coli TB1 pKK540 was digested with Xbal and Pstl, digested with Exonuclease pKK563 PUC18 III, blunt-ended with S1 nuclease, and religated 1.28 kb n.d. E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2664 pKK564 pUC18 III, blunt-ended with S1 nuclease, and religated 2.67 kb (approx.) E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-2791 PKK565 pUC18 III, blunt-ended with S1 nuclease, and religated 2.79 kb (approx.) E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-3002 PKK566 pUC18 III, blunt-ended with S1 nuclease, and religated 3.0 kb (approx.) E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-3084 pKK567 pUC18 III, blunt-ended with S1 nuclease, and religated 3.1 kb (approx.) E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-3244 PKK568 PUC18 III, blunt-ended with S I nuclease, and religated 3.25 kb (approx.) E. coli TB1 pKK540 was digested with Xbal and Pstl, digested with Exonuclease 176 (approx. - PKK569 pUC18 III, blunt-ended with SI nuclease, and religated 3.27 kb 3444 E. coli TB1 pKK540 was digested with Xbal and Pstl, digested with Exonuclease 360 (approx.) • pKK570 pUC18 III, blunt-ended with SI nuclease, and religated 3.09 kb 3444 E. coli TB1 pKK540 was digested with Xbal and Pstl, digested with Exonuclease pKK571 pUC18 III, blunt-ended with SI nuclease, and religated 2.94 kb 502 - 3444 E. coli TB1 pKK540 was digested with Xbal and Pstl, digested with Exonuclease pKK572 pUC18 III, blunt-ended with SI nuclease, and religated 2.79 kb 659 -3444 E. coli TB1 pKK540 was digested with Xbal and Pstl, digested with Exonuclease PKK573 PUC18 III, blunt-ended with SI nuclease, and religated 2.68 kb 768 - 3444 E. coli TB1 Table 9, cont.

pKK540 was digested with Xbal and Pstl, digested with Exonudease 996 (approx.) • pKK574 PUC18 III, blunt-ended with S I nuclease, and religated 2.45 kb 3444 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 1147 (approx.) pKK575 PUC18 III, blunt-ended with S1 nuclease, and religated 2.3 kb -3 4 4 4 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonudease 1262 (approx.) PKK576 p(JC18 III, blunt-ended with S I nuclease, and religated 2.18 kb -3 4 4 4 E. coli T61

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 2263 (approx.) pKK577 PUC18 III, blunt-ended with S I nuclease, and religated 1.18 kb - 3444 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonudease 2462 (approx.) PKK578 PUC18 III, blunt-ended with S I nuclease, and religated 0.98 kb -3 4 4 4 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonudease 2676 (approx.) pKK579 pUC18 III, blunt-ended with S I nuclease, and religated 0.77 kb -3 4 4 4 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 2793 (approx.) PKK580 pUC18 III, blunt-ended with S1 nuclease, and religated 0.65 kb -3 4 4 4 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonudease 2976 (approx.) PKK581 pUC18 III, blunt-ended with SI nuclease, and religated 0.47 kb -3 4 4 4 E. coli TB1 pKKSOI was digested with Xbal and Pstl, digested with Exonudease 1 -1 5 9 PKKS82 PGEM-4Z III, blunt-ended with S1 nuclease, and religated 0.16 kb (approx.) E. coli TB1 pKK540 was digested with Xbal and Pstl, digested with Exonudease 934 (approx.) • PKK583 PGEM-4Z III, blunt-ended with SI nuclease, and religated 2.51 kb 3444 E. coli TB1 Table 9, cont.

pKKS40 was digested with Xbal and Pstl, digested with Exonuclease 3167 (approx.) pKK584 PUC18 III, blunt-ended with S I nuclease, and religated 0.28 kb -3 4 4 4 E. coli TB1

pKK540 was digested with Xbal and Pstl, digested with Exonuclease 3366 (approx.) PKK585 PUC18 III, blunt-ended with St nuclease, and religated 0.08 kb - 3444 E. coli TB1 pKK538 was digested with Xbal and Pstl, digested with Exonuclease 1-3308 pKK586 PUC18 III, blunt-ended with S I nuclease, and religated 3.31 kb (approx.) E. coli TB1 E. coli The 2.5 kb EcoRI/BamHI fragment from pKK501 was ligated into K38/pGP1- pKK587 pT7-7 EcoRI-BamHI-dioested pT7-7 2.5 kb 1-2476 2

The 4.0 kb BamHI fragment of genomic DNA containing hutH from 1-2476 plus S. griseus SKK896 was ligated into the BamHI site of pUC18. The approx. 1.5 kb pKK594 PUC18 direction of the hutH gene is toward the EcoRI site in thepoiylinker. 4.0 kb upstream DNA E. coli TB1

1-2476 plus approx. 1.5 kb pKK595 pUC18 iThe same insert as in pKK594, but in the opposite orientation. 4.0 kb upstream DNA j E. coli TB1 i I SKK821; | j IThe 2.5 kb BamHI fragment from pKK540 was ligated into the Bglll SKK896; site of pi J702. The hutH gene was inserted in the opposite 1SKK910; pKKI100 pi J 702 orientation relative to the mel promoter in the vector. 2.5 kb 1 - 2476 TK24

The 0.16 kb Hindlll/Sstl fragment from pKK573 was ligated into i Hindlll-Sstl-digested plJ2925. The insert in plJ2925 was excised as a Bglll fragment and ligated into the Bglll site of plJ702. The hutH SKK821; PKK1101 DIJ702 coding sequence is in the same direction as the Pmel. 0.16 kb 768 - 923 SKK896 Table 9, cont.

The 0.27 kb Hindlll-Sstl fragment from pKK572 was manipulated as pKKI102 PU702 described for pKK1101. 0.27 kb 659 • 923 SKK821

The 0.42 kb Hindlll-Smal fragment from pKK571 was filled in with Klenow fragment and ligated into the Hindi site of plJ292S. The SKK821; pKKI103 pU702 remainder of the construction was as in pKK1101. 0.42 kb 502-913 SKK896

The 0.16 kb Hindlll-Sstl fragment from pKK573 was ligated into Hindlll-Sstl-digested plJ486. The promoter for hutH was in the TK24; pKK1104 pU486 correct orientation relative to the neo reporter gene of p(J486. 0.16 kb 768-923 SKK821 As in pKK1104, except the hutH promoter was in the incorrect TK24; pKK1105 PIJ487 orientation relative to the neo reporter gene 0.16 kb 768-923 SKK821

The 0.27 kb Hindlll-Sstl fragment from pKK572 was ligated into Hindlll-Sstl-digested plJ486. The hutH promoter was in the correct TK24; pKK1106 plJ486 orientation relative to the neo reporter gene of plJ486 0.27 kb 659-923 SKK821

As with pKK1106, except that the hutH promoter was in the TK24; pKK1107 p(J487 incorrect orientation relative to the neo reporter gene of plJ487 0.27 kb 659-923 SKK821

The 0.38 kb EcoRI-Sstl fragment from pKK500 was ligated into EcoRI-Sstl-digested pU486. The hutH promoter was in the incorrect TK24; pKK1108 pi J 486 orientation relative to the neo reporter gene of plJ486 0.38 kb 544-923 SKK821 As in pKK1106, except that the hutH promoter was in the correct TK24; pKKI109 PIJ487 orientation relative to the neo reporter gene 0.38 kb 544-923 SKK821 As in pKK1100 except that the hutH gene was inserted in the same TK24; PKK1110 PU702 orientation as the mel gene 2.5 kb 1-2476 SKK821 Table 9, cont.

The 4.0 kb BamHI fragment from pKK594, containing hutH from 1-2476 plus SKK896, was ligated into the Bglll site of pU702. The hutH coding approx. 1.5 kb SKK821; pKK1111 PIJ702 sequence was inserted in the direction opposing the mel gene 4.0 kb upstream DNA SKK896 The 0.42 kb Hindlll-Smal fragment from pKK571 was ligated into the Hindi site of pU2925, such that hutH was in the same orientation as lacZ. The insert was excised with Hindlll and BamHI pKKI112 pXE4 and ligated into Hindlll-Bam digested pXE4 0.42 kb 502-913 SKK821 As in pKK1112, except that the hutH promoter was oriented opposite to lacZ in plJ2925 and in the opposite orientation relative pKKI113 pXE4 to the xylE reporter gene of pXE4 0.42 kb 502-913 SKK821

The 0.27 kb Hindlll-Sstl fragment from pKK572 was ligated to Hindlll-Sstl-digested plJ2925, excised by digestion with Hindlll and Bglll, and ligated to Hindlll-BamHI digested pXE4. The hutH pKKI114 pXE4 promoter was oriented in the same direction as the xylE gene 0.27 kb 659-923 SKK821

The 0.16 kb Hindlll-Sstl fragment from pKK573 was ligated to Hindlll-Sstl-digested plJ2925, excised by digestion with Hindlll and Bglll, and ligated to Hindlll-BamHI digested pXE4. The hutH pKKI115 pXE4 promoter was oriented in the same direction as the xylE gene. 0.16 kb 768-923 SKK821 Appondix C

Isolation of Hut' Mutants

162 Mutagenesis and Mutant Isolation

The procedures for isolation of histidine utilization mutants (Hut ) derived from 5. griseus SKK832 were as described previously (Kroening and Kendrick, 1989) with some modifications. Approximately 5X10° spores of SKK832 were suspended in 4.5 ml of 0.05

M malic acid (pH 8.0) containing 1 mg/ml N-methyl-N’-nitro-N-nitrosoguanidine (MNNG;

Sigma). The spore suspension was incubated at 30°C for 30 min with shaking. The extent of killing was approximately 98.5% under these conditions. The mutagenized spores were centrifuged and washed five times with sterile distilled HsO, plated on SpM, and incubated at 30°C for 5 days. The sporulated colonies on SpM were replicated to glucose-histidine minimal medium and glucose-ammonia minimal medium supplemented with 0.1 mM histidine as needed for the auxotrophs and incubated at 26° or 34°C. Hut' mutants were identified as those that grew only on the glucose-ammonia minimal medium. Heat-sensitive mutants grew on glucose-histidine medium only at the lower temperature, and cold-sensitive mutants grew on giucose-histidine medium only at the higher temperature. 164

Table 10. List of Hut' Mutants of S. griseus.

Strain Description of Hut* Phenotype

SKK500 unconditional mutant; deficient in histidase activity

SKK501 unconditional mutant; deficient in formiminoglutaate iminohydrolase activity;

constitutive synthesis of histidase and urocanase

SKK502 unconditioned mutant

SKK503 unconditioned mutemt

SKK504 unconditioned mutant; production of green pigment in glucose-eunmonia

minimal medium

SKK505 unconditioned mutant

SKK506 unconditional mutant

SKK507 cold-sensitive mutant

SKK508 heat-sensitive mutant

SKK509 unconditioned mutant

SKK510 heat-sensitive mutant

SKK511 heat-sensitive mutant 165

Table 11. Activities of the Hut Enzymes in SKK821 and Hut' Mutants of S, griseus.

Specific Activity (nmoi/min/mg protein)" Strain Growth Condition* Histidase Urocanase FIG- lminohydroiasse

SKK821 Glutamate 0.35 <0.1 n.d.d I SKK821 Glutamate + 0.1 mM 1.99 0.74 n.d. L-histidine | SKK832 CAA + 10 mM L-histidine 5.36 2.10 n.d. I SKK832 Glutamate + 0.1 mM 1.57 0.87 n.d. L-histidine I SKK832 Glutamate + Urocanate + 4.28 1.98 n.d. 0.1 mM L-histidine | SKK500 CAA + 10 mM L-histidine <0.1 0.53 n.d. I SKK500 Glutamate + 0.1 mM <0.1 0.20 n.d. L-histidine SKK500 Glutamate + Urocanate + <0.1 0.86 n.d. 0.1 mM L-histidine SKK501 CAA + 10 mM L-histidine 12.3 6.00 <0.1 SKK501 Glutamate + 0.1 mM 5.65 4.89 <0.1 L-histidine SKK501 Glutamate + Urocanate + 6.73 4.97 <0.1 0.1 mM L-histidine

* CAA, complex medium containing casamino acids; glutamate, minimal medium containing L-glutamate as carbon and nitrogen source; when used, urocanate was added as a powder to cultures growing exponentially in glutamate minimal medium, and the cultures were incubated for an additional generation (3 hr) before harvesting the cells for preparation of the extract. b Values are the averages of enzyme activities in a single extract assayedin triplicate. c Formiminoglutamate iminohydrolase. d n.d., not determined.