Genetic system for micromonospora

A culture comprising the microorganism Micromonospora echinospora ssp. calichensis DR46 (ATCC-53591), which produces complementing factor, is described and disclosed.

BACKGROUND OF THE INVENTION 
This invention concerns the genetic manipulation of a new Actinomycete 
Micromonospora echinospora ssp. calichensis, that produces the 
antibacterial and anti-tumor agents called LL-E33288 complex. Such 
antibiotic and anti-tumor agents are described in co-pending U.S. patent 
application Ser. No. 009,321, now U.S. Pat. No. 4,970,198, filed Jan. 30, 
1987, which application is a continuation-in-part of co-pending 
application Ser. No. 787,066, filed Oct. 17, 1986, now abandoned, which is 
a continuation-in-part of application Ser. No. 672,031, filed Nov. 16, 
1984 and now abandoned. Various derivative products of the LL-E33288 
complex are described in co-pending applications Ser. Nos. 004,154 and 
004,153, both filed Jan. 30, 1987. The disclosure of all such applications 
is herein incorporated by reference. The antibiotic and antitumor 
LL-E33288 complex is produced during the cultivation under controlled 
conditions of the new strain of Micromonospora echinospora ssp. 
calichensis. This microorganism is maintained in the culture collection of 
the Medical Research Division, American Cyanamid Company, Pearl River, 
N.Y. as culture number LL-E33288. A viable culture of this new 
microorganism has been deposited with the Culture Collection Laboratory, 
Northern Regional Research Center, U.S. Department of Agriculture, Peoria, 
Ill. on Aug. 9, 1984, and has been added to its permanent collection. It 
has been assigned by such depository the strain designation NRRL-15839. 
Access to such culture, during the pendency of application Ser. No. 
009,321, now U.S. Pat. No. 4,970,198 shall be available to one determined 
by the Commissioner of Patents and Trademarks to be entitled thereto under 
37 C.F.R. .sctn.1.14 and 35 U.S.C. .sctn.122, and all restrictions on 
availability to the public of such culture will be irrevocably removed 
upon grant of a patent on application Ser. No. 009,321, now U.S. Pat. No. 
4,970,198. 
THE INVENTION 
We have developed effective methods to mutagenize strain NRRL-15839 and 
screen for derivative strains that produce no detectable product or an 
altered pattern of product. A protoplasting and regeneration regimen has 
been developed that has resulted in the transformation of strain 
NRRL-15839 with a plasmid. Furthermore, we have isolated and analyzed a 
DNA fragment from strain NRRL-15839 that contains promoter activity 
primarily at that stage in the life cycle of NRRL-15839 that the LL-E33288 
complex is produced. By utilizing these genetic and biochemical advances, 
we can produce novel derivatives of strain NRRL-15839 that have improved 
characteristics (i.e., production of compounds with reduced toxicity), and 
we can enhance product yield. Because yields of the iodo-LL-E33288 complex 
were higher than the bromo-LL-E33288 complex, iodo-LL-E33288 complex was 
used exclusively in our work.

DETAILED DESCRIPTION OF THE INVENTION 
Strain NRRL-15839 grows as a multi-cellular form called mycelia. The 
multi-cellular nature of growing cultures is a problem for mutagenesis; 
cells containing recessive mutations will not be detected in a cluster of 
wild type cells. One must resort to mutagenesis of spores, a non-growing 
unicellular form of the organism, or mutagenesis of growing cells that are 
then subsequently treated by protoplasting (digesting the cell wall) or 
sonicating (mechanically shearing the cell wall). By preparing protoplasts 
or sonicated mycelia, unicellular or near-unicellular genetic units are 
artificially generated. We describe effective protocols involving the 
mutagenesis of fragmented mycelia of strain NRRL-15839 with 
nitrosoguanidine and with ultra-violet light. 
In order to transfer recombinant DNA into the producing organism, it was 
essential to develop a protoplasting-regeneration system, since 
protoplasts will generally take up exogenous DNA, whereas growing mycelia 
or non-growing spores will not. We have included all the details of these 
procedures, because protoplasting and regenerating appear to be delicate 
procedures. We also include details of the procedure for transferring 
genetic material between strains by protoplast fusion. 
In order to clone genes into strain NRRL-15839, it is necessary to develop 
vectors that can replicate in the organism. A derivative of the plasmid 
pIJ486, a Streptomyces plasmid (J. M. Ward et al., Molecular and General 
Genetics, 1986, pp. 468-478), containing a promoter from strain NRRL-15839 
for the kanamycin resistance gene, was used to transform strain 
NRRL-15839. We will also investigate the utility of other plasmids from 
Micromonospora and from Streptomyces. Endogenous plasmids were not 
observed in strain NRRL-15839. Large plasmids might not be detected, 
however. 
The approach we have taken to manipulate the biosynthetic pathway is 
similar in some respects to the study of tylosin production in 
Streptomyces fradiae, or erythromycin production in Streptomyces erythreus 
[Seno, E. T. and C. R. Hutchinson. 1986. in "The Bacteria", Volume IX, (S. 
W. Queener, L. C. Day, editors; Academic Press Inc. New York) pp. 
231-279]. Those studies involved the isolation of mutants blocked in 
antibiotic production, and the cloning of genes involved in antibiotic 
biosynthesis. Mutants that are blocked specifically in the production of 
LL-E33288 complex can be useful in establishing the biosynthetic pathway 
of the drug. Cloning genes involved in LL-E33288 complex biosynthesis can 
be of great value in increasing yields of drug, and in producing novel 
derivatives. 
We have isolated mutants blocked in LL-E33288 complex production. Some 
pairs of blocked mutants can co-synthesize active product. One mutant of a 
pair secretes an intermediate that the other mutant can convert to 
LL-E33288 complex. By isolating and identifying intermediates made by the 
secreting blocked mutants, we can determine precursor components of the 
LL-E33288 complex molecules. By altering the component or intermediate 
that is fed to the converting strain, we can potentially generate novel 
derivatives of LL-E33288 complex by "bioconversion" (Shier, W. T., K. L. 
Jr. Rinehart, and D. Gottlieb. 1969. Proc. Nat. Acad. Sci. 63: 198-204). 
In the case of LL-E33288 complex, bioconversion is a more feasible way to 
produce novel products than chemical modification of LL-E33288 complex, 
due to the complicated and unstable structure of the various components of 
the LL-E33288 complex. 
Another means of characterizing blocked mutants involves feeding cultures 
known components of the LL-E33288 complex. One such component is the 
pseudoaglycone, a degradation product of LL-E33288 complex that lacks the 
rhamnose moiety and an amino sugar. The loss of the amino sugar results in 
more than a 100-fold reduction in biological activity. The ability of 
blocked mutants to convert the pseudoaglycone to LL-E33288 complex 
indicates that they are incapable of carrying out a reaction essential for 
the formation of pseudoaglycone due to a genetic defect. By altering a 
fragment such as the pseudoaglycone it is again possible to produce novel 
products by bioconversion. 
Blocked mutants can also serve as recipients of plasmid DNA in a 
transformation. It is possible to clone the biosynthetic genes by 
complementing blocked mutants with plasmid DNA carrying wild type DNA 
inserts. It is also possible to clone the biosynthetic genes into another 
organism. Streptomyces lividans is a well-characterized host which 
supports the replication of several useful cloning vectors [Hopwood, D. 
A., et. al. 1985. Genetic Manipulation of Streptomyces; A Laboratory 
Manual. (The John Innes Foundation, Norwich, England)]. We can prepare a 
library of Micromonospora DNA fragments ligated into plasmid DNA, 
transform S. lividans, and screen for transformants that synthesize 
LL-E33288 complex. It may prove impossible, however, to clone, and 
express, all the biosynthetic genes on a single DNA fragment. Production 
of LL-E33288 complex in a foreign host might also be lethal. It might be 
more feasible to clone a part of the pathway, by identifying genes coding 
for precursor molecules, that a blocked mutant converts to LL-E33288 
complex. Other means of cloning biosynthetic genes into S. lividans or E. 
coli include selecting resistance to LL-E33288 complex, since drug 
resistance genes may be linked to biosynthetic genes. DNA probes from 
genes known to be involved in polyketiie biosynthesis (Malpartida, R. et. 
al. 1986. Abstr. Fifth Internat. Symp. on the Genet. of Industrial 
Microorganisms. p. 163) may also be used to identify genes with similar 
functions in strain NRRL-15839. 
If the biosynthetic genes cannot be identified directly, we can screen for 
transcripts that are present only at the time of the life cycle when the 
LL-E33288 complex is synthesized; such transcripts may code for enzymes of 
the LL-E33288 complex biosynthetic pathway. A variation of this approach 
involves searching for promoters active only at the time of the 
biosynthesis of LL-E33288 complex, utilizing promoter probe plasmids. 
Finally it may be possible to identify a protein involved in LL-E33288 
complex biosynthesis, and by sequencing the protein, we could design our 
own probe for screening DNA containing biosynthetic genes. 
Cloning and expressing such genes presents the possibility of elucidating 
the pathway, and eventually of manipulating it to our advantage. By 
cloning and expressing a gene coding for the rate-limiting enzyme of the 
LL-E33288 complex biosynthetic pathway, for example, we could increase the 
yield of product. By transferring such genes to other microorganisms, it 
might be possible to produce novel forms of LL-E33288 complex, as has been 
reported for a derivative of actinorhodin within the genera Streptomyces 
(Hopwood, D. A., et. al. 1985. Nature 314: 642-644). It would be an 
important advance, if by cloning and transferring the LL-E33288 complex 
biosynthetic genes, an organism was able to produce a new form of drug 
that retained the anti-tumor function but was less toxic. 
An important aspect of our plan is the ability to express genes at the 
proper time in the life cycle of the organism. It is possible, for 
example, that the expression of the LL-E33288 complex biosynthetic genes 
during the exponential phase might be lethal. One way to regulate 
expression of cloned genes is at the level of transcription, by utilizing 
a DNA sequence preceding a gene or genes that is only active as a promoter 
at the time that LL-E33288 complex is also made. The isolation of such 
promoters is therefore an important step in the establishment of an 
expression system in Micromonospora or Streptomyces. 
A sensitive biological induction assay (BIA), involving the induction of 
.beta.-galactosidase in response to DNA damage in an engineered strain of 
E. coli, and the thin layer chromatography system which separates 
different components of LL-E33288 complex, were utilized to detect 
LL-E33288 complex activity. These systems were essential for the screening 
and characterization of large numbers of mutagenized cells for those with 
altered properties. 
Mutagenesis of Strain NRRL-15839 
In order to isolate interesting mutants it was essential to mutagenize 
strain NRRL-15839. A common mutagenesis regimen for Actinomycetes is to 
treat spores with agents such as UV or nitrosoguanidine (Delic, V., D. A. 
Hopwood, and E. J. Friend. 1970. Mut. Res. 9: 167-182). We mutagenized a 
growing culture of mycelium, and not spores, with these agents. Generally 
mutagenesis with N-nitro-N-methyl-N-nitrosoguanidine (NTG), which acts 
most strongly at DNA replication forks, is enhanced in growing cells 
[Miller, J. H. 1972 Experiments in Molecular Genetics. (Cold Spring Harbor 
Laboratories, Cold Spring Harbor, N.Y.)]. To insure detection of mutants 
carrying recessive lesions, mycelia were sonicated (mechanically sheared) 
after mutagenesis and outgrowth, to provide single genetic units. 
Culturing strain NRRL-15839 for Mutagenesis with NTG 
The growth medium for mutagenesis was GER (Kim, K. S., and D. Y. Ryu. 1983. 
Enz. Mic. Technol. 5: 273-280), a rich medium supporting relatively rapid 
growth of NRRL-15839, containing 100 mM TES 
(N-tris[hydroxymethyl]methyl-2-aminoethane sulfonic acid) adjusted to pH 
7.6. The buffer was added to maintain a relatively high pH on addition of 
NTG, which is a more effective mutagen near pH 8 (Delic, V., D. A. 
Hopwood, and E. J. Friend. 1970. Mut Res. 9: 167-182). During exponential 
phase, before orange pigment production and before mycelial clumping, cell 
density could be monitored with a Klett-Summerson colorimeter. The culture 
of strain NRRL-15839 was grown in a baffled Klett flask, since aeration 
improves growth, and the culture also contained 3 glass beads (4 mm) to 
break up mycelial masses. 
A frozen seed culture of strain NRRL-15839 was added to a 25 ml culture of 
GER medium. Cells were incubated at 32.degree. C. in a New Brunswick G76 
shaking water bath set at 300 rpm. The initial inoculum was 14 Klett Units 
(green filter). After about twenty-eight hours the cells had grown to 118 
Klett Units, or mid-log phase. Cells were harvested by centrifugation and 
resuspended in 10 ml containing 1 mg/ml NTG, 70% GER, and 100 mM TES at pH 
7.6. The culture was incubated in the water bath, and aliquots of culture 
were centrifuged, washed 3 times with GER medium, and grown out by 
inoculating onto two GER plates, and one slant each of 65-15 medium.sup.1 
or yeast dextrose medium.sup.2. 
FNT .sup.1 64-15 medium g/l: glucose 10 g dextrin 20 g yeast extract 5 g 
N.sup.Z Amine A 5 g; CaCO.sub.3 (Mississippi Line) 1 agar 15 g pH 6.7-7.0 
- made up with tap water .sup.2 Yeast Dextrose Medium g/l yeast extract 10 
g glucose 10 g agar 15 g Aph 6.8 - made up with tap water 
A 2 ml sample of the culture was removed at 2.8' and centrifuged in two 
microcentrifuge tubes. The sample was washed three times to remove NTG. 
Other samples were removed from the culture at 35', 91', and 164'. 
Outgrowth was necessary to allow for expression of recessive mutations 
among the many cells of a mycelial clump. We inoculated the slants, since 
they support development of spores, which are single cells. Mutagenized 
spores were eventually used to screen for blocked mutants. Because spores 
took weeks to form, we also grew out the mutagenized samples on 2 GER 
plates as well. After several bacterial generations of growth on GER 
plates a mutant would presumably divide to form a small, genetically 
homogenous cluster within the mycelium. To obtain fragments of mycelia 
that were single genetic units, 1 ml of such mycelia were resuspended in 
GER broth in a microcentrifuge tube jacketed with ice, was sonicated for 
14-25 seconds (MSE Soniprep 150 sonicator, amplitude 16 microns), and 
mycelia were reduced to 2-3 cell size, as determined by microscopic 
observation. Measurements of colony forming units were also helpful in 
optimizing the sonication time; sonication produces more units to form 
colonies, but also kills some fragments. The number of colony forming 
units as a function of sonication time was determined; the 14-25 second 
range was beyond the time maximum colony forming units. 
Mycelia from the 2.8' and 35' samples were grown out for 4 days, and 
mycelial fragments were prepared and inoculated onto GER plates after 
diluting in 10.3% sucrose. The remainder of sonicated or unsonicated cells 
was diluted with glycerol to 20% and stored at -70.degree. C. After 4 days 
of incubation at 30.degree. C., clones were replicated with toothpicks 
onto minimal medium plates or GER plates. While obtaining valuable 
auxotrophic derivatives of strain NRRL-15839, we could also measure the 
mutagenesis frequency from these data. The minimal medium used initially 
was 73-3 with added phosphate, consisting of, per liter, 20 g sucrose, 1 g 
ammonium sulfate, 0.1 g FeSO.sub.4.7H.sub.2 O, 0.2 g MgSO.sub.4.7H.sub.2 
O, 5 g CaCO.sub.3 (Mississippi Lime), 0.05 g K.sub.2 HPO.sub.4, and 20 g 
agar, adjusted to pH 7.5. We later found it was easier to test clones in a 
modified minimal medium (Aux medium), which substitutes 1/10 volume of TES 
1M pH 7.5, and 7.37 g CaCl.sub.2.2H.sub.2 O for Mississippi Lime, which 
confers an opaque grey color to the medium that obscures colonies. Our 
results were more definitive when Noble agar (Oxoid), which was free of 
nutrient contamination, was substituted for Difco agar. 
For the third NTG time point, the outgrowing cells were allowed to incubate 
for 8 days, until colonies were a reasonable size. The 91' treatment with 
NTG resulted in a long recovery time for the culture. Since there was so 
much killing (see below), we replicated colonies directly from the plate 
of outgrown mycelia, and then collected and sonicated the plate culture as 
described above. It seeded likely that for at least some colonies, only 
one cell would have survived from the mycelial cluster that was plated. 
Only small discrete colonies were picked. It was possible that the 
population picked directly from the plate was enriched for mutants that 
did rot grow as well as the wild type on GER plates, and therefore were 
smaller, and remained discrete during the 8 day incubation. After 
individual colonies were picked, the plate culture was harvested and 
sonicated as outlined above. 
Results of NTG Mutagenesis 
The frequency of mutagenesis with NTG generally correlates with killing 
[Miller, J. H. 1972. Experiments in Molecular Genetics. (Cold Spring 
Harbor Laboratories, Cold Spring Harbor, N.Y.)]. It was obvious that a 
dramatic increase in killing occurred with increasing time of NTG 
treatment. After 4 days of growth at 30.degree. C., cells plated from the 
2.8' sample formed a lawn on GER plates; about 5000 colonies grew on the 
plates containing the sample harvested after 35' NTG treatment, about 300 
colonies from the 91' sample, and just 3 colonies for the 164' sample. 
Preliminary analysis of the auxotrophic screen also suggested a very 
successful mutagenesis. After replicating and retesting clones on plates 
containing minimal or GER medium, mutants that failed to grow on minimal 
medium were tested on a series of plates in order to determine their 
nutritional requirement (Davis, R. W., D. Botstein, and J. R. Roth. 1980. 
Advanced Bacterial Genetics, Cold Spring Harbor Laboratory, Cold Spring 
Harbor, N.Y.). Auxotrophs from the various time points are listed below. 
______________________________________ 
Time Nutritional requirements 
of NTG Proportion (number obtained 
treatment of mutants in parenthesis) 
______________________________________ 
2.8' 4/349 1.1% aromatic amino acids (Aro) 
(2), tryptophan (1), 
histidine (1) 
35' 55/765 7.2% Aro (44), uracil + arginine 
(4), adenine (1), trypto- 
phan (1), undefined (5) 
91' 94/400 23.5% Aro (66), tryptophan (9), 
undefined (2), untested 
(15) 
.sup. 91'.sup.3 
12/100 12.0% Aro (11), lysine (1) 
______________________________________ 
FNT .sup.3 replicated directly, prior to sonication 
Results of the mutagenesis showed a high mutation frequency which increased 
with time of NTG treatment. We could define most mutants, consistent with 
the selection of single mutations. Several useful auxotrophic strains were 
obtained, consisting of six classes. We did, however, observe a pronounced 
clustering of mutations. The more highly mutagenized, the narrower was the 
spectrum of mutations observed. Mutations affecting biosynthesis of the 
aromatic amino acids (Aro) were by far the predominant class. It is known 
that NTG can have mutational "hotspots", especially near the origin of DNA 
replication (Guerola, N., J. L. and E. Cerda-Olmedo. 1971. Nature 230: 
122). We were concerned that the high proportion of Aro mutants reflected 
such as skewing in the distribution of mutations. We therefore used 
ultraviolet light (UV) as a second way to induce genetic lesions. 
Mutagenesis with UV Light 
The protocol was designed to minimize the shielding of cells from UV light, 
which would attenuate its mutagenic effect. Cells were resuspended in a 
clear solution that was shallow enough (depth less than 2 mm) to allow all 
cells to be exposed. To overcome shading of cells by other cells within a 
mycelium, we sonicated to break up mycelial clumps to the size of 2-3 
cells. The alternatives were mutagenesis of spores Hopwood, D. A., et. al. 
1985. Genetic Manipulation of Streptomyces; A Laboratory Manual. (The John 
Innes Foundation, Norwich, England); Delic, V., D. A. Hopwood, and E. J. 
Friend. 1970. Mut. Res. 9: 167-182], or of protoplasts. After mutagenesis, 
the culture was grown out for several cell divisions to allow mutants 
containing recessive lesions to form small cell clusters within mycelia. 
The outgrown culture was again sonicated to produce single genetic units, 
and tested for auxotrophy. 
About 5.times.10.sup.9 cells from a growing culture of strain NRRL-15839 in 
GER medium were washed in 10.3% sucrose, sonicated as described above, 
resuspended in 10 ml of 20% glycerol in a Petri dish, and exposed to 
2.4j/m.sup.2 /sec of UV light (256 nm). At 60, 90, and 120 seconds, 2.8 ml 
of cells were taken from the cell suspension, and cells were diluted and 
inoculated onto GER plates for outgrowth. The unused portion of UV-treated 
cells were stored at -70.degree. C. Other variations of the protocol 
included UV mutagenesis of unsonicated cells. Cells grown in GER medium 
containing 0.15% glycine were also treated with UV light, since cells 
growing in the presence of glycine were observed to contain smaller 
mycelia (perhaps because of the effect of glycine on the cell wall). After 
four days colony forming units were determined, as shown below. 
______________________________________ 
Growth Medium: 
Sonication GER GER + glycine 
prior to UV: 
yes no yes no 
______________________________________ 
Colony Forming 
Units/ml 
0 UV 2.5 .times. 10.sup.8 
3.5 .times. 10.sup.7 
4 .times. 10.sup.7 
3 .times. 10.sup.7 
60 sec 2.9 .times. 10.sup.5 
4 .times. 10.sup.6 
2 .times. 10.sup.5 
4 .times. 10.sup.6 
UV 
90 sec 8 .times. 10.sup.3 
1 .times. 10.sup.6 
1 .times. 10.sup.4 
2.3 .times. 10.sup.5 
UV 
120 sec 2 .times. 10.sup.2 
1 .times. 10.sup.5 
1.6 .times. 10.sup.4 
1.5 .times. 10.sup.4 
UV 
______________________________________ 
A survival curve is shown in FIG. 1. For sonicated cells a logarithmic 
relationship was observed for survival as a function of the time of 
exposure to UV light, and killing was very extensive. For mycelia that 
were not sonicated prior to treatment with UV light, killing was less 
extensive, probably because of shading. For early time points survival of 
unsonicated mycelia may be over estimated; if even one cell within a 
mycelial cluster survives, then no killing is detected. 
Because killing by UV light is often correlated with mutagenesis [Miller, 
J. H. 1972. Experiments in Molecular Genetics. (Cold Spring Harbor 
Laboratories, Cold Spring Harbor, N.Y.)], these populations were likely to 
contain mutants. Plate cultures for each regimen were harvested by 
scraping outgrown mycelia into GER medium. For cell suspensions that were 
sonicated prior to UV treatment, cells exposed to UV for 90 seconds were 
used; unfragmented mycelia treated with UV for 120 seconds were harvested. 
In all cases outgrown mycelia were fragmented by sonication. After four 
days cf incubation on GER plates, colonies were screened for growth by 
toothpicking onto minimal agar plates and GER plates. The following 
proportions of auxotrophs were observed for the four mutagenized cultures: 
growth in GER and no sonication prior to UV treatment, 0/370 (less than 
0.3% auxotrophs), growth in GER with sonication, 3/285 (1% auxotrophs), 
growth in GER+glycine and no sonication 0/388 (less than 0.3% auxotrophs), 
and growth in GER+glycine with sonication, 1/312 (0.3% auxotrophs). 
Since sonication enhanced both killing and mutagenesis by UV light, we 
screened the metagenized culture of cells grown in GER, and sonicated 
prior to treatment with UV. We observed a total of 38 auxotrophs of 1820 
screened, or 2.1%. Auxogeny revealed 33 cysteine or methionine requirers, 
3 histidine requirers, 1 threonine requirer, and undefined mutant. Again 
we observed clustering of mutants, this time requirers of organic sulfur 
(cysteine or methionine). 
Summary of Auxotrophs and Pigment Mutants 
We grew up auxotrophs representing every class in liquid Aux medium with 
and without supplements and confirmed all nutritional requirements. Fresh 
colonies from GER plates were washed and resuspended in 5 ml of broth and 
incubated at 28.degree. C., using a roller drum for good aeration. Taken 
together the two mutagenesis procedures resulted in 8 different classes of 
nutritional requirers. The only overlap of auxotrophs between UV- and NTG- 
induced mutants was histidine requirers. 
The uses of auxotrophs include the following. Two classes (Aro, and 
uracil-arginine requirers) were used in a protoplast fusion experiment 
described below. Auxotrophic alleles are of value in assuring that a given 
strain or colony is marked, and distinguishable from contaminants. It may 
prove useful to clone a wild-type gene that compensates for an auxotrophic 
mutation on a plasmid; the uptake and maintenance of the plasmid could be 
selected for by growing transformants on minimal medium. The mutants 
requiring organic sulfur may prove useful for labeling the LL-E33288 
complex, because each LL-E33288 complex component contains 4 sulfur atoms. 
A broad spectrum of mutants would have suggested that any non-lethal mutant 
could be found in our mutagenized populations. It is unclear why certain 
auxotrophs predominated, such as Aro mutants for NTG-treated cells. The 
aro region could be a mutational hotspot site for NTG. It is also possible 
that aro mutations confer a selective advantage during NTG treatment or 
outgrowth. Alternatively the aro region might be highly mutagenic in 
strain NRRL-15839, as the argG locus is in Streptomyces (J. Altenbuchner 
et al., 1984, Molecular and General Genetics, 195 pp 134-138). It is also 
possible that we were counterselecting for classes of auxotrophs. At least 
for Aro, tryptophan, histidine, threonine, and lysine auxotrophs, cells 
grew up faster in Aux medium containing only essential supplements, as 
opposed to Aux medium containing all supplements. Perhaps nonessential 
amino acids interfere with the uptake of the essential amino acid, 
inhibiting growth. The threonine and histidine requirers grew markedly 
poorly on GER medium compared to the wild type. In any case it seemed 
reasonable to test both the NTG- and UV-treated cultures for other 
desirable mutants, such as blocked mutants. 
One other class of mutants that was observed from NTG-treated cells were 
pigment mutants. Wild type colonies of strain NRRL-15839 make an orange 
color after incubation for several days on GER plates. Several clones were 
observed from NTG-treated cells that no longer produced orange pigment. 
Some mutant colonies were white and some yellow. Our initial interest in 
such mutants stemmed from the observation that pigment production in wild 
type cultures occurred at about the time that LL-E33288 complex was 
produced. Further experimentation showed, however, that there was no 
intimate connection between the ability to make pigment and to produce 
drug, since these pigment mutants maintained the capacity to produce 
LL-E33288 complex. These mutants could be useful as recipients in a 
plasmid transformation that restored the capacity to make pigment, due to 
insertion of wild type Micromonospora DNA. Since pigment production is 
temporally controlled, the isolation of the regulatory sites from such 
genes could be useful in establishing a regulated expression system. 
Isolation of Mutants Blocked in LL-E33288 Complex Production 
In order to isolate blocked mutants, we had to miniaturize the fermentation 
conditions so that many clones could be quickly screened. Colonies were 
inoculated directly into 1.5 ml of 73-3I medium.sup.4, using 24 well 
plates (Corning). Each well contained one glass bead to prevent clumping. 
For a quick screen, 3 or 10 .mu.l of culture broth was spotted on a BIA 
plate, which had been inoculated with the E. coli tester strain. 
BIA-active material was identified from wild type cultures within three 
days. Because inocula were not uniform, cultures were incubated for a full 
week to avoid artifacts. 
______________________________________ 
.sup.4 73-3I Medium 
g/l 
______________________________________ 
Sucrose 20.0 
FeSO.sub.4.7H.sub.2 O 
0.1 
MgSO.sub.4.7H.sub.2 O 
0.2 
CaCO.sub.3 (Miss. Lime) 
2.5 
Marcor Peptone 2.0 
Molasses 5.0 
KI 0.1 
made up with tap water 
______________________________________ 
In order to observe specific components of LL-E33288 complex, the 
extraction process also had to be streamlined. For those cultures that 
were positive for BIA-active material by the quick screen, 0.8 ml of 
culture was mixed with 325 .mu.l of an acetone/ethyl acetate mix (8 
volumes acetone/5 volumes ethyl acetate) in a microcentrifuge tube, and 
vortexed well for 2' using a Multi-tube Vortexer (Scientific Manufacturing 
Industries). After centrifuging, 10 .mu.l was spotted onto a TLC 
(Kieselgel 60 F254 from EM Science), and chromatographed as described, 
except just for 20'. Enough active material could be extracted within the 
small volume of the organic phase to detect LL-E33288 complex components, 
after TLC, on a BIA plate. To store the mutants, a 60 .mu.l sample of all 
cultures was added to 40 .mu.l of 50% glycerol, in a well of a 96 well 
plate, and frozen at -70.degree. C. 
By screening clones for BIA activity, several potentially interesting 
nonproducing derivatives were obtained (FIG. 2). We wanted quickly to 
eliminate mutants that failed to produce LL-E33288 complex for extraneous 
reasons related to poor growth or pleiotropic mutations affecting 
secondary metabolism. We had already determined that orange pigment 
production was not intimately associated with LL-E33288 complex 
production, since pigment mutants made BIA-active product. Pigment 
production occurs in the wild type, however, concurrently with LL-E33288 
complex. Those nonproducers that failed to grow to confluence, and/or 
failed to make orange pigment, were discarded. We isolated 10 nonproducing 
mutants that grew to confluence of 240 clones tested, derived from cells 
mutagenized with NTG for 35' and sonicated. Six nonproducers were obtained 
from 240 clones tested, of cells mutagenized with NTG for 35' and then 
sporulated, and 1 of 480 mutagenized with UV. 
Isolation of Mutants Producing an Altered Profile of LL-E33288 Complex 
Components 
The clones that produced BIA-active material were tested for LL-E33288 
complex components by screening with TLC plates as outlined above. A TLC 
of a collection of mutants displaying an altered profile of LL-E33288 
complex components is shown in FIG. 3. One mutant of strain NRRL-15839, 
called DR414, produces only one visible component co-migrating with the 
delta component from NRRL-15839. Another (DR1122) makes no beta component, 
which is the major component produced by strain NRRL-15839. Several 
(DR823, DR1310, DR1523) produce considerably more of faster migrating 
components than strain NRRL-15839. Four of these mutants are from NTG 
mutagenized cells that were sonicated after outgrowth, and the others from 
NTG mutagenized cells that were sporulated. We do not know whether these 
mutants produce novel derivatives of LL-E33288 complex, or make more of 
minor components that are also produced by strain NRRL-15839. If novel 
products are made by one or more mutants, then they could exhibit the 
kinds of characteristics (i.e. reduced toxicity) that would be very 
valuable. If they emphasize a particular component, such as delta for 
DR414, then they could prove useful in isolating this component. 
Co-Synthesis of LL-E33288 Complex by Blocked Mutants 
In order to demonstrate that a given mutant is blocked specifically in the 
LL-E33283 complex pathway, we assayed for product from mixed cultures. If 
two mutants could co-synthesize BIA-active material, then one mutant might 
provide a precursor that overcomes the defect of the second mutant. The 
diagram below is an example of a linear biosynthetic pathway: 
##STR1## 
Mutant DR46 might contain a defect in enzyme 2, and therefore fail to make 
active product. Suppose mutant DR43 contains a lesion in enzyme 1, an 
earlier step in the pathway. If strain DR46 produces intermediate B and 
secretes it into the growth medium, then strain DR43 might take up 
intermediate B and convert it to the final product, since DR43 contains 
enzymes 2, 3 and 4. By taking up compound B, DR43 bypasses its genetically 
defective gene product (enzyme 1). 
The co-synthesis of BIA-active material by two mutant strains does not 
unequivocally demonstrate that each is blocked specifically in the 
LL-E33288 complex biosynthetic pathway. The mutants might actually contain 
nutritional deficiencies, for example, although most classes of auxotrophs 
make at least some product. (Only the adenine requirer and two classes of 
undefined mutants failed to produce BIA-active material of auxotrophs 
tested.) Another possibility is that one mutant fails to produce an 
essential sugar component of LL-E33288 complex. Such a defect would not 
show up as an auxotroph, but the defect could be alleviated by 
supplementing the growth medium with the sugar, rather than a compound 
specific to LL-E33288 complex. However, some mutants that are blocked 
specifically in the LL-E33288 complex pathway could fail to co-synthesize 
if they produce an unstable intermediate, or one that is not secreted or 
taken up. 
We prepared inocula of each mutant by growing to saturation in GER medium, 
washing, and storing 5x concentrated culture in 20% glycerol at 
-70.degree. C. Then 5 .mu.l of a given mutant were inoculated with 5 .mu.l 
of another mutant, or 10 .mu.l of a single mutant were inoculated into a 
well of 73-3I production medium in a 24 well plate. After one week 
cultures were spotted on BIA plates to detect LL-E33288 complex. An 
example of the co-synthesis of mutant DR43 is shown in FIG. 4. Whereas 
strain DR43 grown alone produces no BIA-active material, strain DR43 grown 
with several other mutant strains co-synthesized the LL-E33288 complex. A 
summary of the co-synthesis experiments is shown in FIG. 5. If two mutants 
co-synthesize, they are in distinct complementation groups; if they fail 
to co-synthesize, they may contain mutations affecting the same step in 
the pathway, or they could affect different functions, but fail to 
complement for the reasons given above. 
The results of the co-synthesis showed that every nonproducer 
co-synthesized with at least 2 other mutants. Among the 17 mutants, we 
observed 11 complementation groups, based on the ability of mutants to 
co-synthesize, or based on a distinct pattern of co-synthesis with other 
mutants. One would expect, for such complicated molecules as present in 
LL-E33288 complex, that at least 40 biochemical steps would be involved. 
Eventually we focused on interesting mutants. Some mutants proved to be 
leaky (DR91, DR58, DR112, DR194), and were not followed up because the 
small amount of active material produced by a leaky mutant might have 
interfered with subsequent experiments. In addition DR172 was not examined 
extensively because it had a nutritional requirement. Four strains were 
eliminated from immediate attention because they were of the same large 
complementation group, and we could not distinguish them easily. 
Co-synthesis experiments in liquid culture cannot determine which mutant of 
a pair is secreting a compound, and which mutant is converting the 
compound to active product. In order to distinguish secreters and 
converters, cells were spotted on production medium in a semi-solid 
matrix, containing 0.5% low melt agarose (Seakem), so that maximum 
diffusion was permitted. Mutant cultures were spotted in pairs adjacent to 
each other. After one week of growth, each spot was harvested by peeling 
colonies off of the agar surface, and cells were treated with 20 .mu.l of 
acetone-ethyl acetate (1/1 vol/vol). Three .mu.l of the ethyl acetate 
phase was spotted onto a BIA plate. The mutant of a pair that produced the 
stronger signal was assumed to be the converter and the other strain the 
secreter. Since LL-E33288 diffuses in the agar, results are not 
definitive. Not all pairs on plates co-synthesized detectable material. 
Still we were able to make preliminary determinations of the following 
pairs of mutants: mutants: 
______________________________________ 
Secreter Converter 
______________________________________ 
DR210 DR43 
DR118 DR43 
DR58 DR43 
DR1712 DR43 
DR1316 DR43 
DR46 DR43 
DR43 DR1510 
DR123 DR46 
DR58 DR46 
DR1712 DR46 
DR1316 DR46 
DR123 DR1316 
DR118 DR1510 
______________________________________ 
Assuming that converting strains are blocked earlier than secreting 
strains, a genetic order can be established for the biosynthetic pathway. 
A tentative map of the LL-E33288 complex pathway, ordering mutants 
according to their block, follows. 
##STR2## 
It became clear that mutant DP43 was a particularly good candidate to 
examine in detail. It co-synthesized with several different mutants, and 
experiments suggested that mutant DR43 was usually the converting strain. 
Therefore DR43 probably contains a lesion early in the pathway. The 
clearest delineation on plates was observed when DR43 (converter and DR46 
(secreter) were co-grown. 
A viable culture of mutant strain DR46 is maintained by the Medical 
Research Division American Cyanamid Company, Pearl River, N.Y. 
Additionally, a viable culture of mutant strain DR46 has been deposited 
with the American Type Culture Collection, 12301 Parklawn Drive, 
Rockville, Md. 20852 on Mar. 6, 1987, and has been added to its permanent 
collection. It has been assigned by such depository the strain designation 
ATCC-53591. Access to such culture, during the pendency of the instant 
application, shall be available to one determined by the Commissioner of 
Patents and Trademarks to be entitled thereto, under 37 C.F.R. 1.14 and 35 
U.S.C. 122, and all restrictions on availability to the public of such 
culture will be irrevocably removed upon grant of a patent on the instant 
application. 
Characterization of an Intermediate of LL-E33288 Complex Biosynthesis 
Since mutant DR46 appeared to secrete an intermediate that mutant DR43 
converted to BIA-active material, it was likely that at least some of the 
intermediate would be found in the supernatant of a culture of strain 
DR46. Then it would be possible to add back a cell-free supplement from 
strain DR46, allowing strain DR43 to produce active material. The success 
of such an experiment depends on the stability of the intermediate; if it 
is very unstable, then it may be necessary to grow strains DR43 and DR46 
together to detect co-synthesis. 
The putative intermediate might be found in the supernatant of a culture of 
strain DR46. Alternatively it might be predominantly inside of cells. We 
decided to supplement cultures of strain DR43 with different components of 
a strain DR46 culture in order to distinguish these possibilities. Because 
the time of secretion of the intermediate was not known, a mixture of five 
timed cultures of strain DR46 was prepared. The five 50 ml cultures were 
harvested after 1, 2, 3, 4 and 6 days of growth in 73-3I production 
medium. A mixture of supernatants and cell pellets was prepared. The cell 
pellet mixture was resuspended in 10 ml H.sub.2 O and sonicated for a 
total of 17' (15 seconds sonicating per minute, jacketed with ice, MSE 
Soniprep 150, 15 microns). Few cells remained intact. The sonicate was 
centrifuged to remove debris, and filter-sterilized. The precipitate was 
extracted with ethyl acetate, since LL-E33288 complex is mostly found in 
an ethyl acetate extraction, and is not very soluble in water. The ethyl 
acetate extract was dried and resuspended in ethanol. 
Strain DR43 cultures were inoculated in 24 well plates, and after 1, 2 or 3 
days the components of the strain DR46 mixed culture were added. After 4 
days of growth, BIA-activity was tested by spotting cultures. (To maintain 
a constant volume when large volumes of the strain DR46 culture 
supernatant were added to a well, an equal volume of the strain DR43 
culture was centrifuged, and the pellet was added back.) The supernatant 
of the culture of strain DR46 contained the putative intermediate 
("complementing factor"). 150 .mu.l of supernatant of strain DR46 
contained sufficient complementing material to be detected, following 
conversion by strain DR43, to BIA-active material. 750 .mu.l of 
supernatant from strain DR46 cultures supplemented strain DR43 at least as 
well as the co-synthesis of DR43 and DR46. The equivalent of 1.7 ml of 
cell sonicate supplemented about as well as 150 .mu.l of supernatant, 
indicating that the intermediate was predominantly in the culture 
supernatant, and not inside the cell. The ethyl acetate extract from the 
equivalent of 8 ml of strain DR46 cells (10 .mu.l of concentrated extract) 
did not detectably supplement. 
Since the complementing factor produced by strain DR46 was found primarily 
in the culture supernatant, we concentrated our efforts there. 1.3 ml of 
supernatant from strain DR46 cultures incubated 1 or 2 days, and 130 .mu.l 
of supernatant from strain DR46 cultures incubated 3, 4 and 6 days, were 
added back to cultures of strain DR43 after two days of growth in wells. 
On day 4 samples were spotted on a BIA plate to determine if complementing 
factor was in the supernatant (FIG. 6). In order to observe the BIA 
response accurately, 3 .mu.l samples of culture undiluted, diluted 1/3, 
and diluted 1/10 were applied to the BIA plate. No complementing and 130 
.mu.l of supernatant from strain DR46 cultures incubated factor was 
detected from the strain DR46 supernatant from day 1, and a barely 
detectable amount of complementing factor by day 2. The concentration of 
complementing factor in the supernatant increased substantially by day 3, 
and peaked by day 4. The appearance of complementing factor in the 
supernatant of the strain DR46 culture parallels the appearance of 
LL-E33288 complex made in the wild type. The complementing complementing 
factor is synthesized in the life cycle after the end of growth, as is the 
case for LL-E33288 complex in the wild type (see below). The temporal 
coincidence is consistent with the conception that complementing factor is 
a specific precursor of LL-E33288 complex, and not, for example, a sugar 
that is synthesized during growth and stationary phase. Furthermore, the 
supernatant from S. lividans TK54 grown in 73-3I medium (with added 
histidine and leucine) does not provide complementing factor to strain 
DR43. 
The chemical properties of complementing factor also indicate that it is 
not a simple sugar, but rather a precursor with hydrophobic and 
hydrophilic domains. The complementing factor did not partition into the 
organic phase after ethyl acetate addition, but about 1/2 the 
complementing factor entered the secondary butanol phase when an equal 
volume was added to supernatant. About 1/3 of the complementing factor was 
found in n-butanol when it was mixed with supernatant. 
We have developed a process that an be used to isolate complementing factor 
from large cultures of strain DR46. The complementing factor adsorbs to 
ambersorb XAD-2 beads (Rohm and Haas). It is eluted almost completely with 
80% methanol. The process has been scaled up to enable the isolation of 
complementing factor from a 300 l fermenter. The culture of strain DR46 is 
filtered, and the supernatant of the fermentation is applied to a 30 l 
column of XAD-2 beads. Most of the complementing factor is retained on the 
column, and then eluted with a 50% water-methanol mix. After 
lyophilization, complementing factor has been further purified by 
reverse-phase column chromatography, using an LH20 or a C18 matrix. 
When strains DR43 and DR46 were co-grown, only the beta component of 
LL-E33288 complex was made (see FIG. 7). The same result was observed when 
a culture of strain DR43 was supplemented with complementing factor. Based 
on these observations and the chemical properties of complementing factor, 
we suspect that complementing factor contains sugars, as well as a benzene 
ring (contributing a hydrophobic domain). 
Conversion of Pseudoaglycone to LL-E33288 complex 
The pseudoaglycone fragment of LL-E33288 lacks a rhamnose derivative and an 
amino sugar. The loss of the amino sugar results in more than a 100-fold 
loss of biological activity. Purified pseudoaglycone was added to 73-3I 
cultures of three blocked mutants, strains DR43, DR46 and DR1316, after 60 
hours of cultivation. In each case the blocked mutants were capable of 
converting pseudoaglycone to LL-E33288 complex, as indicated in FIG. 7. 
Thus each strain retained the ability to make the amino sugar derivatives 
and rhamnose derivative, and to bind them to the pseudoaglycone fragment 
(or a part of the pseudoaglycone). For each mutant, pseudoaglycone 
bypassed, or overcame, the genetic block. 
Protoplasting, Regeneration, and Genetic Transfer 
Protoplasting is generally achieved by digesting the cell wall with 
lysozyme. The addition of glycine to the growth medium often facilitates 
protoplasting of Actinomycetes, probably due to incorporation of glycine 
into peptidoglycan of the cell wall. (Examples of protoplasting procedures 
have been collated in reference [Hopwood, D. A., et. al. 1985. Genetic 
Manipulation of Streptomyces; A Laboratory Manual. (The John Innes 
Foundation, Norwich, England.)] 
In order to protoplast strain NRRL-15839, 250-500 .mu.l of an overnight of 
5x concentrated cells, stored in 20% glycerol at -70.degree. C., were 
inoculated into 50 ml of GER medium [per 1, 3 g beef extract, 5 g 
tryptone, 1 g dextrose, 24 g soluble starch and 5 g yeast extract, pH 7.6 
(Kim, K. S., and D. Y. Ryu. 1983. Enz. Mic. Technol. 5: 273-280] 
containing 0.15% glycine and 20 mM CaCl.sub.2, at 28.degree. C. Cells were 
grown for 48 to 60 hours, and the incubation was continued until about 8 
hours after the appearance of orange pigment. Since vigorous aeration was 
found to be essential for good protoplasting, cells were grown in 250 ml 
baffled flasks on a floor shaker set at 200 rpm. The addition of 3 glass 
beads (about 4 mm diameter) prevented clumping, and aided in the growth 
and the protoplasting of cultures. 
Cells were harvested by centrifuging at 3200 rpm for 10'. The pellet was 
washed with 50 ml of 10.3% sucrose and centrifuged again. The pellet was 
resuspended in 4 ml of protoplasting buffer containing mg/ml lysozyme 
(either Calbiochem or Sigma). Protoplasting buffer was similar to L buffer 
[Hopwood, D. A., et. al. 1985. Genetic Manipulation of Streptomyces; A 
Laboratory Manual. (The John Innes Foundation, Norwich, England)], except 
the pH was 7.6. A modified form of P buffer (Kim, K. S., and D. Y. Ryu. 
1983. Enz. Mic. Technol. 5: 273-280) was also employed; MgCl.sub.2 was 25 
mM and CaCl.sub.2 was to 50 mM, and again the pH was 7.6. Cells were 
incubated for one-two hours at 30.degree. C. The extent of protoplasting 
was monitored in the phase contrast microscope. Since mycelia may inhibit 
the regeneration of protoplasts, it was advantageous to wait until the 
vast majority of the lysozyme-treated culture had protoplasted. We 
generally observed more than 95% protoplasting; if fewer than 80% of cells 
were protoplasts, the culture was not used. If the time of protoplasting 
exceeded 2 hours, we did not observe effective regeneration. In all cases 
after protoplasting, the final concentration of MgCl.sub.2 and CaCl.sub.2 
was adjusted to 25 mM and 50 mM respectively, and the volume was brought 
up to 3 ml in our modified P buffer. In order to separate protoplasts from 
mycelia, to enhance the frequency of regeneration, we filtered through 
cotton as described [Hopwood, D. A., et. al. 1985. Genetic Manipulation of 
Streptomyces; A Laboratory Manual. (The John Innes Foundation, Norwich, 
England)], after first rinsing the preparation with modified P buffer. 
Alternatively we filtered through Schleicher and Schuell Spartan filters 
with 5 micron pore size, after rinsing with modified P buffer. 
Protoplasts were regenerated by inoculating them onto regeneration medium 
(as described in Kim, K. S., and D. Y. Ryu. 1983. Enz. Mic. Technol. 5: 
273-280) except that the optimal concentration of sucrose was found to be 
0.15M. The efficiency of regeneration was improved by overlaying 
protoplasts in soft agar. Protoplasts regenerated to form visible colonies 
after incubation at 30.degree. C. in 10 days to 2 weeks. 
Several modifications of the regeneration conditions were attempted, but 
without success. Succinate and glycerol were ineffective as osmotic 
stabilizers, because mycelia could not tolerate them at high 
concentrations. The presence of alternative carbon sources would probably 
not alter carbon utilization in regeneration medium, since sucrose, a 
preferred carbon source, was required as the osmotic stabilizer. Other 
nitrogen sources, yeast extract at 0.5%, or NZ amine at 0.5%, were 
substituted for asparagine. We observed only mucoid colonies, albeit at a 
reasonable frequency, compared to our regeneration medium containing 
asparagine. The colonies, in the microscope, appeared to contain L forms 
and not mycelia. Furthermore the colonies would not replicate onto media 
lacking osmotic stabilizer. Apparently regeneration of protoplasts is 
enhanced in medium containing a growth-rate limiting nitrogen source. We 
also attempted to regenerate protoplasts on GER medium containing sucrose 
as an osmotic stabilizer, and some added minerals. Again mucoid colonies 
grew up at a reasonable frequency, but they appeared also to be L forms. 
The following is a typical successful experiment. Cells were harvested and 
treated with lysozyme as described above. After 1.5 hours at 30.degree. 
C., about 95% of cells were protoplasts, estimated by microscopic 
observation. The concentration of protoplasts was determined to be 
2.9.times.10.sup.9 /ml, using a Petroff-Hauser counting chamber. The 
culture was gently filtered through a 5 micron filter. Only protoplasts, 
and about 1% of small mycelial fragments, were observed in the microscope. 
The titer of protoplasts was 2.35.times.10.sup.9 /ml. Protoplasts were 
diluted in modified P buffer and plated on regeneration medium. The 
culture was also plated on regeneration medium that lacked sucrose, in 
order to measure cells that hadn't been protoplasted, since protoplasts 
would lyse in the absence of osmotic stabilizer. After 3 days a few 
colonies were visible for the less dilute inocula, for plates with or 
without 0.15M sucrose. These colonies presumably represent cells that were 
not protoplasted. After 1 week more colonies were visible due to 
protoplast regeneration. After 11 days almost all the surviving 
protoplasts formed visible colonies. 
______________________________________ 
0.2 M % 
Sucrose Dilution Colonies Colonies/ml 
Survival 
______________________________________ 
- 5 .times. 10.sup.-5 
394 8 .times. 10.sup.6 
0.3% 
- 5 .times. 10.sup.-7 
12 2 .times. 10.sup.7 
0.9% 
+ 5 .times. 10.sup.-5 
3240 6.5 .times. 10.sup.7 
2.8% 
+ 5 .times. 10.sup.-7 
50 1 .times. 10.sup.8 
4.2% 
______________________________________ 
The addition of sucrose enhanced survival of the protoplast culture, even 
though 0.2M sucrose inhibits growth of mycelia. Therefore, the enhanced 
survival in the medium containing sucrose is due to the regeneration of 
protoplasts. Our frequency of regeneration of from 1-4% is reasonable for 
a transformation system. 
In order to demonstrate genetic exchange between derivatives of strain 
NRRL-15839, we attempted a protoplast fusion between two auxotrophs. 
Strain DR226 requires uracil and arginine for growth, while strain DR146 
requires aromatic amino acids, para-amino benzoic acid, and para-hydroxy 
benzoic acid. Both mutants may contain a single lesion affecting one 
biochemical reaction (production of carbamoll phosphate for the 
uracil-arginine requirer, and of shikimic acid pathway for the aromatic 
amino acid requirer). By fusing protoplasts of the two mutants, we hoped 
to observe the regeneration of wild type cells. 
Cells were protoplasted as described above. 100 .mu.l of DR226 was mixed in 
a microcentrifuge tube with 100 .mu.l of DR146. After centrifugation for 
10 seconds, the supernatant was poured off, and protoplasts were gently 
resuspended in the remaining drop of supernatant. 0.8 ml of a 50% 
polyethylene glycol solution (PEG 1000, Baker) was added in modified P 
buffer as described. In order to select for prototrophs, a minimal 
regeneration medium lacking 0.01% casamino acids, was used, since the 
regeneration frequency was found to be about the same with or without 
casamino acids. 50 .mu.l of fused protoplasts were overlaid cnto minimal 
regeneration agar plates containing no osmotic stabilizer, or 0.2M 
sucrose. We also plated an equal volume of mycelia before protoplasting as 
a control. After four weeks the following colonies were observed. 
______________________________________ 
0 M sucrose 
0.2 M sucrose 
______________________________________ 
DR146 mycelia 0 
DR226 mycelia 1 
DR146 and DR226 protoplasts 
1 30 
______________________________________ 
The number of prototrophs among mycelia is practically 0, whereas after 
protoplasting the two cultures and fusing, a small but significant number 
of prototrophs regenerated (the maximum regeneration frequency observed is 
4%). The fusion of auxotrophs to prototrophy demonstrates the ability to 
transfer genes within two derivatives of NRRL-15839. It is now possible to 
transfer mutations from one strain to another. 
Constructing a Plasmid for Transforming Strain NRRL-15839 
We did not detect endogenous plasmids within NRRL-15839. Micromonospora is 
related to Streptomyces, and several Streptomyces plasmids are availalle. 
We therefore decided to construct a derivative of a well-characterized 
plasmid from Streptomyces, with the hope that it would also replicate in 
strain NRRL-15839. 
The Streptomyces promoter probe plasmid pIJ486 contains the thiostrepton 
resistance gene on a multi-copy plasmid. Since its replicon derives from 
pIJ101, which has a broad host range within Streptomyces, pIJ486 seemed to 
be a good starting point for a Micromonospora plasmid. The thiostrepton 
resistance gene was not useful, since strain LL-E33288 is partially 
resistant to at least 500 .mu.g/ml thiostrepton in GER or minimal agar 
medium. pIJ486, however, also contains another potentially useful gene, 
the kanamycin resistance gene from the transposon tn5, called aphII. The 
aphII gene, however, lacks a promotar, and is therefore expressed at very 
low levels. Promoters can be identified by cloning fragments into a 
polylinker upstream of the coding region for aphII, transforming S. 
lividans, and selecting or screening for kanamycin resistance. To 
"activate" the aphII gene, we inserted DNA fragments from strain 
NRRL-15839 for two reasons. A promoter derived from Micromonospora would 
afford the most chance of driving expression of aphII when the plasmid 
derivative is transferred from S. lividans to strain NRRL-15839. Also, a 
DNA sequence derived from Micromonospora could permit an alternative mode 
of maintenance of the plasmid derivative, by homologous recombination, if 
the plasmid replicon failed to function in strain NRRL-15839. 
Plasmid was prepared from S. lividans TK54/pIJ486 by alkaline lysis with 
NaCH-SDS, and neutralization with acidic K acetate as described [Hopwood, 
D. A., et. al. 1985. Genetic Manipulation of Streptomyces; A Laboratory 
Manual. (The John Innes Foundation, Norwich, England.)], and then by CsCl 
density gradient centrifugation [Maniatis, T., E. F. Fritsch, and J. 
Sambrook. 1982. Molecular Cloning. (Cold Spring Harbor Laboratory, Cold 
Spring Harbor, N.Y.)]. The purified plasmid was then digested at the 
unique BamHI site, extracted with phenol, and precipitated with ammonium 
acetate and ethanol. The linear plasmid molecule was treated with 
calf-intestinal alkaline phosphatase to prevent ligation of vector to 
vector ends. The enzyme was inactivated by heat killing and phenol 
extraction. After precipitation with acidified Na acetate and ethanol, the 
vector was resuspended in I.E buffer (10 mM Tris pH 8, 1 mM EDTA), and was 
ready for ligation. All enzymes were from Boehringer Mannheim, and all 
reactions were carried out as recommended by Boehringer Mannheim. 
Micromonospora chromosomal DNA was isolated using lysozyme-EDTA, 
proteinase K, SDS, and phenol-chloroform extraction as described [Hopwood, 
D. A., et. al. 1985 Genetic Manipulation of Streptomyces; A Laboratory 
Manual. (The John Innes Foundation, Norwich, England.)]. After isopropanol 
precipitation the DNA preparation was resuspended in 1 ml TE. A streak of 
the culture revealed that it was pure. 
Chromosomal DNA prepared from strain NRRL-15839 was digested with BamHI and 
Sau3A1, and the fragments were ligated to 0.5 .mu.g of the linearized 
pIJ486 vector. The molar ratio of insert to vector was 3. After ligating 
overnight at 16.degree. C. (Boehringer Mannheim enzyme and methods), the 
DNA was used to transform S. lividans strain TK54 as described. For pIJ486 
ligated to Sau3A1 inserts, 0.25 .mu.g of ligated DNA was added to 
protoplasts of TK54, which were inoculated on an R5 plate and incubated 
for 30 hours at 28.degree. C. The plate was overlayed with 1/10 volume of 
R5 soft agar containing 100 .mu.g/ml of kanamycin. For pIJ486 ligated to 
BamHI inserts, 0.5 .mu.g of ligated DNA was added to protoplasts of TK54, 
and aliquots were inoculated onto two plates. One was overlayed with 
kanamycin as indicated, and the other with 500 .mu.g/ml thiostrepton in 
the overlay. For the latter plate, 71 colonies grew following addition of 
thiostrepton. For the plates overlaid with kanamycin, a lawn of cells grew 
on much of the plate. From patches of plates, single colonies were 
isolated. We observed the following results when colonies were replicated 
onto minimal medium containing the drug indicated. 
__________________________________________________________________________ 
# of Number of Resistant Colonies 
Ligation Colonies 
50 micro g/ml 
2 micro g/ml 
20 micro g/ml 
and Selection 
Replicated 
thiostrepton 
kanamycin 
kanamycin 
Clones 
__________________________________________________________________________ 
Transformation A 
56 54 37 17 pPP4 
BamHI fragments 
selected with 
thiostrepton 
Transformation B 
100 5 28 9 pPP8 
BamHI fragments 
selected with 
kanamycin 
Transformation C 
100 43 45 6 pPP14 
Sau3A1 fragments 
selected with 
kanamycin 
__________________________________________________________________________ 
Many of the clcnes selected directly with kanamycin were artifacts. In 
transformation A, only 5 colonies were resistant to thiostrepton. We 
assumed that thiostrepton-sensitive clones were not true transformants. 
Transformation B was more effective Among 45 putative kanamycin resistant 
transformants, 43 were also resistant to thiostrepton. Of the 56 clones 
selected with thiostrepton (transformation C), 54 proved to be 
thiostrepton resistant when retested, and of those, 34 were at least 
marginally kanamycin resistant, suggesting that most inserts containei 
promoters directing expression of aphII. 
To show that clones resistant to kanamycin and thiostrepton contained 
plasmid, we isolated plasmid from 17 clones that were the most kanamycin 
resistant, judging by colony size on plates containing 20 .mu.g/ml 
kanamycin. Ten were from transformation A, 1 from transformation B, and 6 
from transformation C. Plasmid minipreps were prepared as outlined above 
for plasmid preparations, except that minipreps were scaled down as 
described. Plasmids were observed in each preparation by agarose gel 
electrophoresis. At least 14 of the clones contained plasmid larger than 
the pIJ486 control, suggesting that kanamycin resistance was due to a 
Micromonospora DNA insert in front of the aphII gene. The insert size was 
determined for 5 clones by restriction analysis to be 8.5 Kbp (plasmid 
pPP4), 1.8 Kbp (plasmid pPP8), 0.7 Kbp, 0.7 Kbp, and 0.4 Kbp (plasmid 
pPP14). The 3 clones that were most resistant to kanamycin, carrying 
plasmids pPP4, pPP8, and pPP14, grew on R5 agar medium containing 200 
.mu.g/ml kanamycin and 50 .mu.g/ml thiostrepton, and on minimal plates 
containing 20 .mu.g/ml kanamycin. 
The presence of Micromonospora inserts in these three plasmids was 
confirmed by Southern analysis, as in Schleicher and Schuell (FIG. 8). 
Micromonospora DNA and S. lividans TK54 chromosomal DNA was digested with 
BamHI, applied to a 0.7% agarose gel, and denatured in alkali. DNA was 
transferred to a nitrocellulose filter, which was hybridized with probe of 
purified pPP8 DNA (0.5 .mu.g) labeled with .alpha.-.sup.32 P-dCTP by nick 
translation (NEN nick translation kit). About 10.sup.6 cpm of probe was 
hybridized with the filter at 42.degree. C. The final wash was at 
55.degree. C. at 0.1.times.SSC. Lane one is a control showing the 
hybridization of probe to 150 ng of pPP8, restricted with BamHI. The probe 
hybridizes to two bands; the 1.8 Kbp insert band, and the larger band 
representing linear pIJ486, which is 6.2 Kbp. In lane 2, 2.5 .mu.g of 
Micromonospora DNA, restricted with BamHI, was applied. The 1.8 Kbp band 
is clearly visible, indicating its presence in the Micromonospora 
chromosome. Lane 3, containing S. lividans DNA, gives no signal, 
indicating that the 1.8 Kbp is not derived from the S. lividans genome. 
Similar experiments were performed using pPP4. Two BamHI bands were 
observed when the probe was hybridized to Micromonospora DNA digested with 
BamHI, and no bands with S. lividans DNA, indicating that pPP4 contains 
two BamHI fragments from Micromonospora that add up to 8.5 Kbp. When pPP14 
was used as probe, again a band was observed only when Micromonospora DNA, 
cut with BamHI, was probed, and not S. lividans DNA. In all three cases, 
therefore, the DNA inserts, activating expression of the aphII gene, were 
derived from strain NRRL-15839. 
Plasmids pPP4, pPP8 and pPP14 were placed in host S. lividans TK54 and 
deposited with the American Type Culture Collection, 12301 Parklawn Drive, 
Rockville, Md. 20852 on Mar. 6, 1987, and have been added to its permanent 
collection. pPP4 has been assigned by such depository the strain 
designation the strain designation ATCC- 67,343, pPP8 has been assigned by 
such depository the strain designation AICC-67342, and pPP14 has been 
assigned by such depository the strain designation ATCC-67341. Access to 
such cultures, during the pendency of the instant application, shall te 
available to one determined by the Commissioner of Patents and Trademarks 
to be entitled thereto under 37 C.F.R. .sctn.1.14 and 35 U.S.C. .sctn.122, 
and all restrictions on availability to the public of such cultures will 
be irrevocably removed upon grant of a patent on the instant application. 
Transformation of Strain LL-E33288 
Plasmid pPP8 was used to transform protoplasts of NRRL-15839, since it 
conferred resistance to kanamycin and contained a sizable insert (1.8 
Kbp). Our rationale was the following. If the replicon of pIJ486 failed to 
replicate in strain NRRL-15839, then the plasmid could still be maintained 
by homologous recombination into the Micromonospora chromosome. A plasmid 
with a smaller insert (i.e. pPP14) is less likely to undergo 
recombination. Plasmid pPP4 seemed a less attractive choice since its 
insert (8.5 Kbp) dramatically increased the size of the plasmid. If 
Micromonospora contains a restriction system capable of digesting 
unmodified foreign DNA, then it would probably restrict such a large 
plasmid more severely than pPP8. 
Protoplasts of strain NRRL-15839 were prepared as previously described 
above, using 5 micron filters following lysozyme treatment. Protoplasts 
were transformed with pPP8 plasmid DNA, using the protocol similar to that 
used for Streptomyces transformations [Hopwood, D. A., et. al. 1985. 
Genetic Manipulation of Streptomyces; A Laboratory Manual. (The John Innes 
Foundation, Norwich, England.)]. 100 .mu.l, containing 3.4.times.10.sup.8 
protoplasts, was centrifuged for 10 seconds (microcentrifuge), the pellet 
was decanted, and protoplasts were carefully resuspended in the remaining 
drop of supernatant. 18 .mu.l of pPP8 (4 .mu.g) was added, and then 100 
.mu.l of 25% PEG 1000 in T buffer was added. Controls were plated to 
indicate the regeneration frequency, and most of the transformation mix 
was overlayed onto 4 RM plates that had 0.15M sucrose. 
Transformants were selected by overlaying the plates with 150 .mu.g/ml of 
kanamycin in 1/10 volume of soft agar. It was not clear, however, wher to 
add the drug. Transformants might not survive if kanamycin were added too 
early, before drug resistance was well expressed, and/or before fragile 
protoplasts began to regenerate. If kanamycin were added after the optimal 
time, then transformants might not be detected if the plasmid were not 
stably maintained in strain NRRL-15839, or if mycelia inhibited the 
regeneration of protoplasts. We therefore decided to add the kanamycin 
overlay at two times; after one day of incubation at 28.degree. C., and 
after 6 days. Six colonies grew from plates overlaid after 1 day and 127 
colonies when plates were overlaid after 6 days. Colonies were replicated 
on GER plates containing no kanamycin, or 1 or 5 .mu.g/ml kanamycin. The 
17 colonies that grew well in the presence of kanamycin were retested on 
plates with 50, 15 and 5 .mu.g/ml of drug, and their resistance to 
kanamycin suggested that they might contain plasmid. Furthermore, these 17 
colonies appeared to be slightly more resistant to thiostrepton (500 
.mu.g/ml in GER plates); kanamycin-resistant clones grew up in the 
presence of thiostrepton one day before the wild type colonies. 
Plasmid minipreps were made for six clones that grew in the presence of 50 
.mu.g/ml kanamycin, from GER plate cultures containing 5 .mu.g/ml 
kanamycin. Ten .mu.l of a 25 .mu.l plasmid preparation was applied to a 
0.7% agarose gel. Though the chromosomal band in our minipreps was 
prominent, it was difficult to tell whether a plasmid band was present. It 
was conceivable that the plasmid pPP8 was present in our kanamycin 
resistant strains, but at a lower copy number than in S. lividans. It was 
also possible that plasmid sequences were maintained after integration 
into the bacterial chromosome. In order to increase our detection 
capabilities, and to distinguish between these two possibilities, a 
Southern transfer was performed (Schleicher and Schuell method) and the 
parent plasmid, pIJ486, was used as probe. 
The Southern hybridization is shown in FIG. 9. Lanes 1 and 2 contain pPP8 
purified plasmid and a miniprep of S. lividans strain TK54/pPP8 DNA fixed 
to the filter; there are two DNA bands that hybridize to the probe, 
corresponding to the covalently closed circular and nicked plasmid forms. 
An inspection of the stained gel, prior to transfer, indicated the 
position of the chromosomal DNA band, which was clearly above the 
supercoiled plasmid band. In lane 3, the prominent signal from one 
putative Micromonospora transformant is in the same position as covalently 
closed circles of pPP8 from S. lividans. Thus pPP8 replicates autonomously 
in strain NRRL-15839, and is not integrated. 
The miniprep of the Micromonospora plasmid was also restricted with BamHI, 
which should release the 1.8 Kbp insert from the plasmid, resulting in a 
linear form of pIJ486. A Southern of that gel is shown in FIG. 10. Lane 1 
shows a purified plasmid preparation of pPP8 cut with BamHI, and probed 
with labeled pIJ486. Lane 2 shows no signal when DNA from strain 
NRRL-15839 is run in the gel. In lane 3 DNA from the Micromonospora 
miniprep was cut with BamHI before gel electrophoresis. The probe 
hyridized with a band corresponding to pIJ486, confirming the presence of 
plasmid in the transformant. 
Another plasmid miniprep from the Micromonospora transformant was used to 
transform S. lividans TK54 protoplasts to thiostrepton resistance. Three 
thiostrepton resistant colonies were obtained. Plasmid was prepared from 
one clone, and agarose gel electrophoresis revealed a plasmid of the size 
of pPP8. The recovery of a plasmid from Micromonospora that confers 
thiostrepton and kanamycin resistance confirms independently that we 
isolated a Micromonospora transformant. 
Characterization of a DNA Sequence Promoting Transcription in the 
Stationary Phase, Concurrent with LL-E33288 Production 
Our original goal was to define the promoters discussed above that were 
derived from Micromonospora, and to show whether initiation sites used by 
strain NRRL-15839 were similar to those of Streptomyces lividans, a 
relatively well-characterized organism. It became increasingly evident 
that we had one DNA sequence from Micromonospora that had unexpected and 
potentially very useful properties. This DNA fragment contains multiple 
promoters of two types. One promoter is active predominantly in growing 
cells, an activity that was selected for by demanding kanamycin 
resistance. In addition the fragment contains a cluster of 3 promoters 
which are most active when cells have entered stationary phase and the 
drug LL-E33288 complex is produced. We now have the capacity to regulate 
the expression of genes so that they will be expressed at a particular 
time of the life cycle, by juxtaposing a given structural gene downstream 
of the promoter-bearing fragment, or a smaller derivative of it. It may be 
advantageous to express certain genes only during stationary phase, since 
it may be lethal if some genes are expressed during the exponential phase. 
A structural gere, for example, coding for an enzyme that is part of the 
LL-E33288 complex biosynthetic pathway could be placed after our DNA 
sequence, so that the enzyme activity would only appear when drug is made. 
A detailed analysis of promoters from Micromonospora contained on plasmid 
pPP14 is described below. 
Southern analysis had shown that plasmids pPP4, pPP8 and pPP14 all 
contained DNA derived from strain NRRL-15839 inserted in front of the 
aphII gene (FIG. 8). Since the DNA sequences promote expression of the 
aphII gene in S. lividans, we decided to examine their promoter activity 
in Micromonospora, by isolating RNA from strain NRRL-15839 and measuring 
transcription. 
A frozen seed culture of cells was inoculated into 50 ml of GER medium 
(3/100 dilution) and after 60 hours incubation, the culture was mixed with 
frozen medium and centrifuged for 3' at 10000 rpm, and the cell pellet was 
stored at -70.degree. C. until used. To prevent degradation of RNA, the 
cells were rapidly harvested and quick-cooled. Pellets were resuspended in 
guanidinium isothiocyanate solution (Chirgwin, J. M., et. al. 1979. 
Biochem. 18: 5294-5299), and sonicated for a total of 1-2', at 16 micron 
power in the Soniprep 150 (MSE); at least 90% of cells were lysed by 
microscopic observation. RNA was isolated by centrifuging through a CsCl 
cushion (Turpen, T. H., and O. M. Griffith. 1986. Biotechniques 4: 11-15), 
extracted with phenolchloroform, ethanol precipitated, and stored as an 
ethanol precipitate at -20.degree. C., or in water at -70.degree. C. To 
identify the endogenous transcripts coded by the inserts of pPP4, pPP8 
and pPP14, the RNA from strain NRRL-15839 was applied to an agarose gel 
containing formaldehyde [Maniatis, T., E. F. Fritsch, and J. Sambrook. 
1982. Molecular Cloning. (Cold Spring Harbor Laboratory, Cold Spring 
Harbor, N.Y.)], transferred to a filter, and hybridized to labeled pPP4, 
pPP8 or pPP14 (Northern analysis). 
FIG. 11 is the first Northern experiment. Two transcripts, 450 and 375 
bases long, hybridize to the pPP14 probe. Because pPP14 hybridized to two 
distinct transcripts, it appeared that at least two signals involved in 
the regulation of transcription were within the Micromonospora DNA insert. 
The fragment could contain two or more promoters (sites essential for 
initiation of transcription), or a promoter and a 
transcription-termination site. 
In order to precisely identify transcription initiation sites within the 
0.4 kbp Micromonospora DNA insert, we next began a series of S1 nuclease 
transcriptional mapping studies. The 0.4 kbp insert was subcloned into 
pUC19, an E. coli vector, by utilizing the HindIII site of the polylinker, 
and the BamHI site proximal to aphII, and transforming E. coli strain JM83 
(New England Biolabs) to ampicillin resistance. (One BamHI site was 
recreated when the Sau3A1 fragment was ligated into the BamHI site of 
pIJ486.) The resulting plasmid, designated pPEC14, was used for subsequent 
labeling experiments. The fragment was labeled by kinasing the BamHI site 
(Boehringer Mannheim enzyme and reaction conditions), and digested with 
HindIII enzyme, and single-stranded probe was isolated [Maniatis, T., E. 
F. Fritsch, and J. Sambrook. 1982. Molecular Cloning. (Cold Spring Harbor 
Laboratory, Cold Spring Harbor, N.Y.)]. Excess of the probe was hybridized 
with 10 .mu.g of RNA from strain NRRL-15839 or from S. lividans 
TK54/pPP14, and the hybridized samples were treated with S1 nuclease at a 
final concentration of 300 units/ml (Boehringer Mannheim), as outlined 
(Brosius, J., R. L. Cate, R. Perlmutter. 1982. J. Biol. Chem. 257: 
9205-9210). After ethanol precipitation, samples were analyzed by gel 
electrophoresis and autoradiography. 
FIG. 12 shows that RNA from the GER culture of strain NRRL-15839, 
hybridizes and protects the probe from S1 nuclease digestion. Four 
apparent start sites were detected; one start site approximately 150 bp 
from the BamHI end, and a cluster of 3 start sites about 225 bp from the 
BamHI end. Apparently there are multiple start sites for transcription in 
the same direction within this Micromonospora DNA fragment. RNA was also 
prepared from a GER culture of S. lividans TK54/pPP14 and hybridized to 
the probe, ani S1 transcriptional mapping analysis indicated that the 
start sites were the same in the two species. One additional start site 
apparently occurs in S. lividans that is not detectable in Micromonospora, 
about 170 bp from the BamHI site. Because the pattern of transcription was 
so similar in the two species, the approach of selecting for promoter 
activity in S. lividans is valid. 
A map of the 0.4 kbp fragment, showing the transcription start sites as 
well as useful restriction sites is shown in FIG. 13. The spacing between 
the downstream promoter (designated P2) and the 3 upstream promoters 
(designated P1a, P1b and P1c) was about the same as the spacing between 
the RNAs observed with the Northern analysis (FIG. 11). Thus, it seemed 
likely that the 0.4kbp fragment contained several initiation sites for 
transcripts that had a common 3' end. We found no detectable transcript 
when the other strand was labeled and hybridized with S. lividans RNA, 
indicating no promoter activity in the diverging direction. 
To correlate transcription from P1 and P2 with the two bands observed by 
Northern analysis, the following experiment was performed. Derivatives of 
the pPEC14 plasmid were constructed that contained a subfragment of the 
0.4 kbp sequence into pUC19, by utilizing the SstII site in the middle of 
the 0.4 Kbp insert. The upstream portion (containing the P1 cluster) was 
deleted by digesting with PstI and SstII, blunting with T4 DNA polymerase 
(Boehringer Mannheim) [Maniatis, T., E. F. Fritsch, and J. Sambrook. 1982. 
Molecular Cloning. (Cold Spring Harbor Laboratory, Cold Spring Harbor, 
N.Y.)], and ligating to form pPEC2. The downstream portion of the 0.4 kbp 
fragment (containing P2) was deleted by digesting pEC14 with SstII and 
SmaI, and then blunting and ligating to form pPEC1 (FIG. 13). The 
prediction was that pPEC1, containing P1, should hybridize with only the 
larger transcript in a Northern. pPEC2, containing P2, should hybridize to 
both transcripts. If the two transcripts were not overlapping then pPEC1 
and pPEC2 would each hybridize to only one band. In fact, labeled pEC2 
hybridized to both transcripts, and labeled pPEC1 to just the larger 
transcript when the Northern was performed. Thus all the data is 
consistent with overlapping transcripts. 
A high-resolution Northern experiment was done in order to see if the 
larger transcript actually contains multiple bands in the RNA preparation, 
consistent with initiation from P1a, P1b and P1c. The RNA sample from 
Micromonospora was applied to a 5% acrylamide/0.27% bis gel containing 7M 
urea. After electrophoresis, the RNA was electroblotted onto 
nitrocellulose, and probed with labeled pPP14 DNA. FIG. 14 shows that the 
larger transcript hybridizirg to probe pPP14 can be resolved into more 
than one component. Since multiple large transcripts are detectable in the 
RNA preparation, the multiple promoters, P1a, P1b and P1c do not appear to 
be artifacts of S1 nuclease digestion. 
In order to delineate precisely tha locations of the start sites for P1a, 
P1b, P1c and P2, DNA was sequenced from the AvaI site, downstream of P2, 
by the Maxam-Gilbert method (Maxam, A. M., and W. Gilbert. 1980. Methods 
Enzymol. 65: 499-560). For analysis of P1, DNA was sequenced from the 
HpaII/NciI site. DNA was labeled by kinasing, for the non-coding strand, 
and by filling in with DNA polymerase Kleno fragment (Boehringer Mannheim 
enzyme and reaction conditions) for the coding strand. The P1 and P2 start 
sites were defined by lining up the fragment protected from S1 digestion 
with a sequencing gel (FIG. 15). Parts of the sequence of the 0.4 kbp 
fragment were determined by Sanger sequencing (IBI Kit) of plasmids pEC1 
and pEC2. 
Since multiple promoters were used to transcribe the same downstream RNA, 
we decided to examine whether differential utilization of these promoters 
occurred at distinct phases of the growth cycle of Micromonospora. Strain 
NRRL-15839 was grown up in 73-3I medium containing added phosphate 
(K.sub.2 HPO.sub.4 to 0.05%), and growth was monitored by a DNA assay 
using Hoechst 33258 reagent (Calbiochem) as described (McCoy, KW. F., and 
LB. H. Olson. 1985. Appl. Environ. Microbiol. 49: 811-817). (Similar 
growth curves were obtained by measuring protein, utilizing the 
BIO-RAD.TM. assay.) A culture of strain NRRL-15839 was also grown in a 
minimal production medium which we developed (Aux, except K.sub.2 
HPO.sub.4 was to 0.05%, KI was added to 0.01%, and the pH of the TES 
buffer solution was 7.0). 
Growth curves of the culture are shown in FIG. 16. As indicated in the 
figure, BIA activity was not detected until at least 6 hours after the end 
of the exponential growing phase. The Northern analysis of RNA for these 
time points, using labeled pPEC14 as probe, is shown in FIG. 17. The first 
lane contains RNA isolated from exponentially growing cells; very little 
of the larger transcripts starting from P1 was detected, and little of the 
375 base transcript, from P2. As cells were harvested progressively later 
in stationary phase, the larger transcripts from P1 became considerably 
more prominent, whereas the 375 base transcript from P2 diminished 
slightly (lanes 2-5). Interestingly, the prominence of transcripts 
originating from P1 parallels the appearance of LL-E33288 complex in the 
growth medium. We have observed substantially the same results using 
staged RNA from cells growing in minimal production medium (FIG. 17), or 
from S1 nuclease transcriptional mapping studies of staged RNAs. We 
estimate there is at least a ten-fold increase in expression originating 
from P1 during stationary phase, compared to exponential phase. 
The P2 promoter also appears to be subject to regulation. When cells of 
strain NRR-15839 were grown in GER medium, expression of P2 was more 
pronounced during exponential phase than for cells grown in either 73-3I, 
or in minimal production medium (data not shown). During stationary phase 
in GER medium, transcription from P2 increased, whereas transcription from 
P2 diminished for cells grown in produrtion media (FIG. 17). The rich 
nitrogen-containing supplements in GER, compared to production media, may 
account for these differences in expression from P2 in strain NRRL-15839. 
Since the P2 promoter contributes significantly to transcription only 
during exponential phase for strain NRRL-15839 growing in these production 
media, we decided to construct plasmids that contained only P1 or only P2. 
Plasmid pPEC1 was digested with HindIII and EcoRI, and the fragment 
containing P1a, P1b and P1c was inserted into pIJ486, the promoter-probe 
plasmid from Streptomyces, using its HindIII and EcoRI sites, this plasmid 
being designated pSL1. Similarly, a fragment containing P2 was obtained by 
cutting pPEC2 with BamHI and HindIII, and ligating into the BamHI and 
HindIII sites of pIJ486, this plasmid being designated pSL2. By making 
these constructs we coud independently assess promoter activity from P1 
and from P2. We found that insertion of either sub-fragment in front of 
the aphII gene confers kanamycin resistance (up to 40 .mu.g/ml on RM 
plates) to the S. lividans TK54 host. S1 transcriptional mapping analysis 
indicates that the start site of P2 in this construct is identical to the 
P2 start site of pPP14. 
The fact that the insert containing only P2 has significant promoter 
activity eliminates the possibility that the shorter transcript results 
from processing of a transcript originating from P1. The promoter activity 
of P1 in the absence of P2 will be determined to see if the induction 
during stationary phase is still 10 fold. The data indicate that in strain 
NRRL-15839, and in transformants of S. lividans, the P1 promoters are 
temporally regulated. 
Construction and screenino of strain NRRL-15839 genomic DNA library in E. 
coli 
We have constructed two genomic libraries of NRRL-15839 DNA in E. coli. 
Genomic DNA was prepared from strain NRRL-15839 and partially digested 
with Sau3AI restriction endonuclease (Boehringer Mannheim). he digested 
DNA was size fractionated in order to maximize for large inserts in 
cloning (&gt;10 kb). DNA fractionated by sucrose gradient centrifugation was 
ligated into the BamHI site of plasmid pBR322 (New England Biolabs), and 
was used to transform E. coli strain K802 (New England Biolabs) to 
ampicillin resistance. The choice of the E. coli host strain was found to 
be critical to ensure high frequency of transformation. Three thousand 
recombinant clones were obtained, with an average insert size of 14 kbp. 
Alternatively, DNA from strain NRRL-15839 was size fractionated by agarose 
gel electrophoresis, and this DNA was ligated into the BglII site of a 
derivative of plasmid pKK233-2 (Pharmacia), which we modified by insertion 
of a polylinker. This vector contains the inducible trc promoter and 
allows expression of Micromonospora DNA in E. coli. Approximately 3000 
clones containing plasmids with an average insert size 10 kb were 
obtained. 
Hybridization of Micromonospora DNA to Polyketide Probes 
DNA of strain NRRL-15839 was isolated, digested with BamHI restriction 
enzyme, and fractionated on an agarose gel for Southern hybridization 
analysis. The polyketide probes, PIJ2345 and pIJ2346, derived from the act 
I and act III genes of S. coelicolor A3(2) (F. Malpartida et al., 1986 
Molecular and General Genetics, Vol. 205, pp. 66-73), were labeled by 
nick-translation. A BamHI fragment of about 2.5 kbp cross-hybridized with 
the act I probe. 
Physioloqical Studies 
The development of a defined redium that supports the production of 
LL-E33288 complex, provides the opportunity to study the physiologiral 
factors affecting drug production. The favored carbon source for growth of 
strain NRRL-15839 is sucrose or glucose. The favored nitrogen source is 
ammonium. When the minimal medium is changed, and contains alternative 
carbon and nitrogen sources, the production of drug was observed. When the 
concentration of axmonium, in particular, was low, then more drug was 
produced per cell mass. Glutamate (to 1%) was the better nitrogen source 
for drug production when compared to ammonium, proline, or arginine. 
Therefore, the nature of the nitrogen source may play an important role in 
the production of LL-E33288 complex. The nature of the carbon source was 
less important. 
The inventors would like to acknowledge S. Lucania (E. J. Squibb, New 
Brunswick, N.J.) for providing thiostrepton.