I. SLOW ADAPTATION OF UREASE LEVELS IN TOBACCO CELLS CULTURED
ON VARIOUS SOURCES OF NITROGEN

3
II. THE INITIAL ORGANIC PRODUCTS OF ASSIMILATION OF [1 N]AMMONIUM
AND [13N]NITRATE BY TOBACCO CELLS IN CULTURE

BY

Thomas A. Skokut

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY

Department of Botany and Plant Pathology

1978

ABSTRACT

I. SLOW ADAPTATION OF UREASE LEVELS IN TOBACCO CELLS CULTURED
ON VARIOUS SOURCES OF NITROGEN

BY

Thomas A. Skokut

The influence of urea and other sources of nitrogen on the
level of urease in tobacco (N. tabacum L. cv. Xanthi) XD cells
in culture was investigated. Cells cultured on nitrate as the
sole nitrogen source have specific activities of urease that are
only 1/4 to 1/5 of those in cells which have been cultured on
urea for over a period of a year (urea-conditioned cells).

When cells which were previously grown on nitrate are
transferred to urea medium, the activity of urease in these
cells does not increase to the higher level found in urea-
conditioned cells but remains at the lower level normally found
on nitrate grown cells. When such cells are continuously
cultured on urea, it takes at least 40 generations before their
urease activity increases to the higher level. When urea-
conditioned cells which possess the high levels of urease are

transferred into nitrate, ammonium succinate or casein

Thomas A. Skokut

hydrolysate medium, the activities of urease in these cells do
not immediately decrease to the lower levels, but gradually
decrease over several transfers.

Clones isolated from the population of cells grown always
on nitrate have urease activities which do not vary considerably
from the low activity found in uncloned samples of the entire
cell population. From this measured variance, the expected
frequency of high-urease cells in the population was calculated.
The probability that any cells in this population possess a high
level of urease is extremely small. From the measured rate of
doubling of high-urease cells (which is slightly greater than
that of low-urease cells) the time required for selection to
produce a high-urease culture was also calculated. Selection
of a spontaneous variant cell which possesses the high urease
activity or selection of a pre-existing subpopulation of cells
which possess the high urease activity would have required far
more than 40 generations for the urease levels to increase.
These results, plus the observation that clones isolated from a
cell population grown on nitrate and transferred to urea medium
all exhibit a gradual increase in urease, indicate that the rise
in urease activity is based upon an adaptation of all the cells.

The possibility that the mechanism responsible for the slow
adaptation of urease levels may be a multiplication of genes for

urease is discussed.

II. THE INITIAL ORGANIC PRODUCTS OF ASSIMILATIONCE‘[13NIAMMONIUM
AND [13N]NITRATE BY TOBACCO CELLS IN CULTURE

BY

Thomas A. Skokut

Glutamine is the first major organic product of assimilation

13 + .
of NH4 by tobacco cells grown on nitrate, urea, or ammonium
succinate as the sole source of nitrogen, and of 13NO'- by

3

tobacco cells cultured on nitrate (13N05' was used here for the

first time for a study of assimilation of nitrogen). The
percentage of 13N in glutamate, and subsequently in alanine,

increases with increasing times of assimilation. Methionine

sulfoximine inhibits the incorporation of 13W from 13NH;. into

glutamine more extensively than it inhibits the incorporation of
13N into glutamate, with cells grown on any of the three

sources of nitrogen. Azaserine strongly inhibits glutamate

synthesis when 13NH2' is fed to cells cultured on nitrate.

These results indicate that the major route for assimilation

of 13NH;' is the glutamine synthetase—glutamate synthase

pathway, and that glutamate dehydrogenase also plays a

Thomas A. Skokut

role, but a minor one. Methionine sulfoximine inhibits the

. . 3 - .
incorporation of 1 N from l3NO3 into glutamate more strongly

than it inhibits the incorporation of 13N into glutamine,

suggesting that the assimilation of 13NH;- derived from 13N0;'

may be mediated solely by the glutamine synthetase-glutamate

synthase pathway.

ACKNOWLEDGMENTS

I would like to thank my advisors Dr. Philip Filner and
Dr. C. Peter Wblk for the sound scientific advice they gave me
during the course of these investigations. They gave me freedom
to pursue my own scientific interests but were always there when
I needed them. Because of their guidance, being a graduate
student at the Plant Research Laboratory was a stimulating and
rewarding experience.

I would also like to thank the other members of my
guidance committee, Dr. Deborah Delmer, Dr. Norman Good and
Dr. Peter Carlson, for their helpful comments and critical
reading of the manuscript.

I would especially like to acknowledge those persons who
helped with the 13N experiments. Dr. Joseph Thomas, Dr. Jack
Meeks, Dr. WOlfgang Lockau, Dr. Norbert Schilling, Dr. W.-S.
Chien, and Mr. Paul Shaffer all contributed invaluable
collaborative assistance. I would like to also thank Dr. James
Tiedje, Dr. Richard Firestone, Ms. Mary Firestone and Mr. Scott
Smith for assistance in irradiation of the aqueous target, and
in analytical and preparative use of HPLC.

Most of all, I thank my wife Michiyo for her encouragement
and patience.

This research was supported by the U.S. AEC/ERDA/DOE under
contract EY-76-C-02-1338 and by U.S. NSF grants PCM-74—01206 and

78-01684.

LIST OF ABBREVIATIONS

MS Murashige and Skoog medium
TRIS 2-Amino-2(hydroxymethyl)-l,3-propandiol
DTE dithioerythritol

gr.fr.wt. gram fresh weight

MES 2(n-morpholino)ethane sulfonic acid
HPLC high performance liquid chromatography
MSX methionine sulfoximine

iii

TABLE OF CONTENTS

LIST OF TABLES . . . . . . . . . . . . . .

LIST OF FIGUES O O O O O O O O O O O I O 0

INTRODUCTION . . . . . . . . . . . . . . .

LITERATURE REVIEW. . . . . . . . . . . . .

Urea and Ammonium

Growth. . . . . . . . . . . . . . .
Cellular Urea . . . . . . . . . . . .
Occurrence of Urease. . . . . . . . .

Physiological Significance of Urease in

Regulation of Urease. . . . . . . . .
Extraction and Purification of Urease
Assays for Urease Activity. . . . . .
Catalytic Properties of Urease. . . .
Molecular Properties of Urease. . . .
ATP-Dependent Hydrolysis of Urea -

Pathway . . . . . . . . . . . . . .

Nitrogen for Plant

Allophanate

Slow Changes in Phenotypic Expression . . . . . . .

MATERIALS AND METHODS. . . . . . . . . . .

The Tobacco Cell Line and Growth Conditions . . . .

Growth Media. . . . . . . . . . . . .
Harvesting of Cells and Preparation
Assay for Urease. . . . . . . . . . .
Assay for Ammonium Ion. . . . . . . .
Determination of Urea . . . . . . . .
Determination of Protein Content. . .
Cloning . . . . . . . . . . . . . . .

RESULTS 0 O O O O O C C O l O O O I O O I 0

Growth of Cells as the Only Source of
Activity of Urease in Cells Grown for
Generations on Nitrate or Urea. . .

of Extracts . .

Nitrogen. . .
many

Urease Activity of Cells Transferred from Nitrate

Medium to Urea Medium . . . . . . .

Is the Increase in Urease Due to the Selection of

a Spontaneous Variant? . . . . . .

iv

Page
vi

viii

31

31
31
32
33
34
35
36
36

38
38

38
53

66

Page

Is the Increase in Urease Due to the Selection of a

Large Pre-existing Subpopulation of High-Urease

Cells? . . . . . . . . . . . . . . . . . . . . . . . 67
Increase in Urease Activity During Growth on Urea

in Clones Derived from a Population of Cells

Previously Grown on Nitrate. . . . . . . . . . . . . 72
Comparison of Growth Rates of Cultures which Exhibit

the High or Low Levels of Urease and Calculations

for Rise in Urease Due to Growth of High-Urease

Cells. . . . . . . . . . . . . . . . . . . . . . . . 74
Effect of Mixing Nitrate—Conditioned and Urea-

Conditioned Cells on the Rise in Urease Activity . . 78
The Effect of Various Nitrogen Sources on the Activity

of Urease in Urea-Conditioned Cells. . . . . . . . . 80
The Effects of Threonine on the Growth of Cells on

Urea and Other Nitrogen Sources. . . . . . . . . . . 96

Effect of Threonine on Growth of Cells which are

Newly Transferred from Nitrate to Urea or

Ammonium Succinate Medium. . . . . . . . . . . . . . 99
Effect of Tungstate on Growth of Cells . . . . . . . . 104
Search for the Presence of Urea Amidolyase in the

Cultured Tobacco Cells . . . . . . . . . . . . . . . 104
Effect of Nickel and Potassium Citrate on Growth

of Cells . . . . . . . . . . . . . . . . . . . . . . 106

DISCUSSION. 0 o I O o o I o o o o o c o o o o o o o o o o o 110

BIBLIOGRAPHY o o o o o o g Q g o o o . o o o o o o 9 o 9 O o 121

INTRODUCTI ON 0 O O . O O O O I . O O O I O O O O I C 0 C O O 1 3 4

MATERIALS AND METHODS . . . . . . . . . . . . . . . . . . . 139

The Tobacco Cell Line and Growth Conditions. . . . . . 139
Generation ofBNH4+ and 13NO3’ . . . . . . . . . . . . . 139
Assimilation of 13NH4’and 13NOj’ . . . . . . . . . . 142

RESULTS . . . . . . . . . . . . . . . . . . . . . . . . . . 144

Assimilation of 13NH4 . . . . . . . . . . . . . 144
Inhibition of Assimilation of 13NH4+ by Methionine
Sulfoximine and Azaserine. . . . . . . . . . . . . . 150
Pulse-Chase Experiments. . . . . . . . . . . . . . . . 152
Assimilation of 13N05' . . . . . . . . . . . . . . . . 152

DISCUSSION. 0 O O O O O O O O O O C O O O O O I O O O O O Q 162

BIBLIOGRAPHY. . . . . . . . . . . . . . . . . . . . . . . . 167

Table
I.

II.

III.

IV.

VI.
VII.

VIII.

IX.

XI.

II.

LIST OF TABLES

PART I

Variability of activity of urease as measured
with the standard assay. . . . . . . . . . . . .

Effect of extract from cells grown on nitrate
on urease activity of extract from cells grown
on urea o o o o o o o o g o o o o o o o o o o o o o

Urease activity of clones isolated from a
population of cells grown on nitrate and
transferred to urea medium . . . . . . . . . . .

Urease activity of mixtures of urea—conditioned
cells and nitrate-conditioned cells cultured on
urea medium. . . . . . . . . . . . . . . . . . . .

Effect of threonine
on nitrate, urea or

on growth of cells maintained
ammonium succinate . . . . . .

Effect of threonine on urease activity in vitro. .

Effect of threonine on growth of cells transferred
from nitrate medium to nitrate, urea or
ammonium succinate . . . . . . . . . . . . . . . .

Effect of threonine on growth of cells transferred
from urea medium to urea, nitrate or ammonium
succinate medium . . . . . . . . . . . . . . . . .

Effect of tungstate on growth of cells maintained
continuously on nitrate, urea or ammonium
succinate. . . . . . . . . . . . . . . . . . . . .

Test for stimulation of urea hydrolysis in
Presence Of ATP 0 O O O Q C O O I O O O I O I O Q 0

Effects of potassium citrate and NiSO4 on growth
of cells on urea, nitrate or ammonium succinate. .
PART II

Mean fraction of 13N in glutamine and glutamate
after assimilation of 13NH4+for 10 or 15 min . . .

Effect of 1 mM methionine sulfoximine on the
appearance of 13N in glutamine and glutamate in
suspensions of tobacco cells exposed to 13NH3‘
and13N03-ooococ'oooccococooco

vi

48

73

79

98
100

101

103

105

107

109

149

151

Table Page

III. Effect of 1 mM azaserine on the appearance of
13N in glutamine and glutamate in suspensions of
nitrate-grown cells exposed to 13NH3’. . . . . . . . 153

vii

Table Page

III. Effect of 1 mM azaserine on the appearance of
13N in glutamine and glutamate in suspensions of
nitrate-grown cells exposed to 13NH3'. . . . . . . . 153

vii

Figure

1

10

11

12

13

LIST OF FIGURES

PART I

Growth of XD cells on urea as the sole source

of nitrogen. . . . . . . . . . . . . . . .

Specific activities of urease expressed on
fresh weight basis for cells grown on urea
nitrate O O O Q I O O O O O O Q I O O O O 0

Specific activities of urease expressed on
protein basis for cells grown on urea or
nitrate I O O I O C Q C O Q I O I O O O C .

Urease of cells grown on nitrate and urea:
proportionality with time and enzyme
concentration. . . . . . . . . . . . . . .

Dependence of the rate of NHj'formation on

a
or

urea

concentration in extracts of cells grown on urea

or nitrate O O O O O O O O O O O O O O O 0

Difference in the activities of urease in
cultures grown on nitrate or urea over the
period of a year . . . . . . . . . . . . .

The levels of urease in cells transferred from

nitrate to urea medium . . . . . . . . . .

Urea concentration of cells consistently

maintained on urea and of cells newly transferred

into urea medium . . . . . . . . . . . . .

Long Term Experiment No. 1: urease activity of

cells transferred from nitrate to urea medium and

consistently maintained on urea thereafter . . .

Long Term Experiment No. 2: urease activity of

cells transferred from nitrate to urea medium and

consistently maintained on urea thereafter . . .

Urease activities of clones derived from

populations of cells having low and high levels

of urease. . . . . . . . . . . . . . . . .

Differences in growth on urea of high-urease

and low-urease cells . . . . . . . . . . .

Decrease in urease activity in urea-conditioned

cells transferred to nitrate medium. . . .

viii

Page

40

43

45

50

52

55

58

60

63

65

69

76

82

Figure

14

15

16

17

18

Page

Gradual decrease in urease activity in urea-
conditioned cells transferred to nitrate
medium and maintained on nitrate medium. . . . . . 85

Decrease in urease activity in urea-conditioned
cells transferred to ammonium succinate medium . . 88

Gradual decrease in urease activity in urea-

conditioned cells transferred to ammonium

succinate medium and maintained on ammonium

succinate. . . . . . . . . . . . . . . . . . . . . 90

Increase in urease activity when cells
previously grown on ammonium succinate are
transferred back to urea medium. . . . . . . . . . 93

Decrease in urease activity in urea-conditioned
cells transferred to casamino acids medium . . . . 95

PART II

Scan of radioactivity from 13N in electrophoretograms

of organic compounds extracted from nitrate-grown,
cultured tobacco cells with 80% methanol after 600,

120, 30, and 10 s of assimilation . . . . . . . . . 146

Distribution of 13N in organic substances soluble

in 80% methanol after assimilation of 13NH4+ for

10, 30, 120 and 600 s by cultured tobacco cells

grown on nitrate, urea or ammonium succinate as

the sole nitrogen source. . . . . . . . . . . . . . 148

Distribution of 13N in organic substances

extracted with 80% methanol from nitrate-grown,

cultured tobacco cells when assimilation of 13NH4+

for 1 min without supplemental NH4+ was followed

by assimilation in the presence of 10 mM NH4+ for

O, 10, 30 and 60 s. . . . . . . . . . . . . . . . . 155

Distribution of 13N in organic substances soluble

in 80% methanol after assimilation of 13NO3" for

l, 2, 6, and 15 min by cultured tobacco cells

grown on nitrate as the sole nitrogen source. . . . 156

Distribution of 13N in organic substances

extracted with 80% methanol from nitrate-grown,

cultured tobacco cells when assimilation of

13N05' for 5 min was followed by assimilation in

the presence of 10 mM N05' for 0, 2, and 5 min . . 160

ix

I. SLOW ADAPTATION OF UREASE LEVELS IN TOBACCO CELLS CULTURED

ON VARIOUS SOURCES OF NITROGEN

INTRODUCTION

It has been known for years that urea as a sole source of
nitrogen can support plant growth and can be as effective as
nitrate or ammonium. Urea is made utilizable as the result of
the presence of the enzyme urease in plant tissue. Urease
hydrolyzes a molecule of urea to CO2 and two molecules of NH;:
thus making the nitrogen of urea available to the plant in the
form of ammonium ion.

Urease was the first enzyme to be purified in crystalline
form (163), and a great deal is known about its molecular
properties (142). These studies have been possible because the
seeds of the jack bean plant are an excellent source of urease.
However, very little is known about the physiological role of
urease in the plant. It is generally assumed that its main
function is to hydrolyze urea. When urea is the only nitrogen
source available, this is very important. Increases in urease
activity of 2 fold in rice plants (119), 2 to 20 fold in potato
plants (126), and 10 to 20 fold in soybean cells in culture (137)
occur as a response to the presence of urea. There is some
evidence that ammonium may repress urease synthesis in rice
plants (119) and soybean cells in culture (137).

l

This study was initiated to find out more about the
regulation of urease in higher plant tissue. A tobacco cell
culture system was used to find answers to the following
questions:

1) Is the enzyme induced by its substrate urea, and does
this induction behave similarly to the induction of nitrate
and nitrite reductase and derepression of ATP sulfurylase which
have previously been studied in these cells?

2) Are the levels of activity detected in the cells affected
by different sources of nitrogen?

3) Do the different cells of a particular cell population

have markedly different levels of the enzyme?

LITERATURE REVIEW

Urea and Ammonium as Sources of Nitrogen for Plant Growth

 

The major forms of nitrogen applied to the soil are nitrate,
ammonium and urea. It is generally assumed that nitrate is the
major form used by plants because ammonium and urea are converted
to nitrate by soil bacteria in a process known as nitrification.
Since nitrate is readily utilized by plants and its assimilation
is closely regulated (52), nitrification can therefore be
considered beneficial. However, nitrification has some
disadvantages to the plant. It results in a product which is
more easily leached through the negatively charged soil
particles, thereby making it more difficult for the soil to
retain its nitrogen. Also, there is a higher susceptibility to
nitrogen loss from the soil through volatilization since
denitrification cannot occur unless the nitrogen in the soil is
first oxidized to nitrate (1). There are many factors which
affect nitrification. Bactericidal agents and fungicides have
a definite effect of inhibiting nitrification by blocking the
growth of the nitrifying organisms (1). There is a decrease in
nitrification with decreasing temperatures (62). When the pH of

the soil is greater than 7.2, the first step in nitrification

(oxidation of ammonium to nitrite) is unaffected, whereas the
oxidation of nitrite to nitrate is greatly suppressed.
Consequently when urea or ammonium is applied to neutral or
alkaline soils in sufficient quantity substantial amounts of
nitrite can accumulate (30,33). Other factors affecting
nitrification are the type of soil, the time of application of
fertilizer, the moisture content of the soil and the aeration of
the soil (1). Because there are various instances when
nitrification is inhibited, at these times ammonium and urea
would be present in the soil as sources of nitrogen for the
plant.

Ammonium and urea are directly utilized without prior
conversion to nitrate as sources of nitrogen by plants. A
summary of numerous investigations which compared the growth
under sterile conditions of various plant species on nitrate,
ammonium and urea (19) concluded that all three compounds were
acceptable sources of nitrogen. Generally the lowest yields
were obtained with ammonium. This might have been due to the
decrease in pH which occurs when plants take up ammonium. It
has been shown that plants can utilize ammonium more readily if
the pH is not allowed to decrease (6,11). Urea supported more
growth than ammonium and in some instances plants grew better
on urea than on nitrate (19). It was also concluded that
growth on urea resulted from the direct uptake of urea and not
ammonium ions which might be present due to the breakdown of

urea (19). A study of urea uptake in roots of rice plants

demonstrated that urea was absorbed faster than either ammonium
or nitrate (89). The use of urea as a fertilizer has been
beneficial in numerous agricultural situations, particularly
when urea is applied as a foliar spray. Foliar feeding of urea
to pineapple plants (184), citrus (110) and apple (60,61) trees,
and banana plants (65) has proved to be very effective and in
some cases more effective than soil fertilization (22). Most of
these studies concluded that growth on urea resulted from the
formation of ammonium in the leaf by the action of urease.
Ammonium and urea have also been shown to be effective
sources of nitrogen for growth of plant tissue cultures.
Ammonium is usually given in combination with nitrate (127)
because most plant tissue cultures will not grow on nitrate
unless a reduced source of nitrogen is also present (67). It
has been shown that cells grown on ammonium and nitrate
utilized the ammonium first, and then the nitrate (10). In
some instances nitrate may be used as the only source of
nitrogen (141,66). It had been difficult to grow plant cell
cultures with ammonium as the sole source of nitrogen until
Gamborg and Shyluk (68) demonstrated that soybean cells could
grow in suspension culture with ammonium alone if a Kreb's cycle
dicarboxylic acid was also present in the medium. Others since
then have used this combination to grow tobacco, tomato and
carrot cell suspension cultures on ammonium (12,40,133,137).
Behrend and Mateles (13), using succinate in combination with

ammonium, showed that the succinate had no effect on the rate of

uptake of ammonium and that succinate was also taken up by the
cells. These authors suggested that the dicarboxylic acid
was needed as an additional source of carbon skeletons for
amino acid synthesis. It has recently been suggested that the
role of the dicarboxylic acid is to limit the pH changes in
the medium brought about by growth on ammonium (41). This
study showed that suspension cultures of carrot would grow on
ammonium as a sole source of nitrogen in the absence of any
exogenous Kreb's cycle acid when the pH of the medium was

controlled by continuous titration with KOH or KHCO Urea

3.
supports growth of plant tissue cultures without specific
supplemental additions to the medium. Cells of tobacco, tomato,

soybean, sycamore, Paul's scarlet rose and carrot have been

grown on urea alone (12,81,96,137,201).

Cellular Urea

 

There are many reports in the older literature of the
occurrence of free urea in plants. However, most of these are
suspect because ureides present in the plant extract were
probably broken down to urea during sample preparation (19).
Although it is presently thought that the cellular concentration
of urea in most plant tissues is generally low or undetectable
(19), there are specific instances when urea may accumulate.

Urea is detected in plant tissue when it it given as a
source of nitrogen. It has been found in rice stems (89),

xylem sap of young apple trees (19) and above ground potato

plant tissue (126) as a result of direct absorption by the
roots, and in leaf tissue which has been sprayed with a urea
solution (65,110).

Urea can be produced endogenously in the plant as a
breakdown product of several nitrogenous compounds. It is a
product of purine degradation in which the following sequence
of reactions is believed to occur: purine + uric acid +
allantoin + allantoic acid + urea + glyoxylate (141). Allantoic
acid accumulating plants degrade adenine-8-14C to urea and
glyoxylate (140), and seedlings of jack pine readily convert
guanine-8-14C to urea (134). Since allantoin has been shown to
be a major molecule for transport of nitrogen in some plants (9,
120) it is conceivable that urea is formed in these plants as a
degradation product of allantoin. Urea can also arise from
the breakdown of arginine. There is considerable evidence that
the reactions of the Krebs-Henseleit cycle (109) exist in
plants. The evidence lies with the detection of all the
intermediates of the cycle and of the enzymes involved (21,97,
105,107,108,153,154,160). In animals this cycle is directed to
the production of urea. Although all of the reactions of the
cycle are present in plants, they do not necessarily function
as a cycle: e.g., in develOping seedlings the enzyme responsible
for the last reaction of the cycle, arginase, independently
produces urea by its action on arginine (37,97,202). Seeds
which contain a high amount of canavanine as a storage compound

for nitrogen have been shown to degrade this compound to

canaline and urea (149,178,190).

Occurrence of Urease

 

The first report of the hydrolysis of urea by an enzyme
called urease was by Musculus in 1876 (128). He separated
bacteria from urine by filtration on filter paper, dried the
paper, impregnated it with litmus and showed that ammonia was
produced from urea because it caused the indicator to turn blue.
Urease was first detected in higher plants by Takeuchi in 1909
(170) when he reported that the soybean seed contained a very
active enzyme which split urea into ammonium carbonate. A few
years later Kiesel (102) reported that urease was present in
many higher plants and in 1916 Mateer and Marshall (117)
discovered that jack bean (Canavalfizensifbnmim seeds exhibited
sixteen times the urease activity of soybean seeds. Sumner
(165) estimated that 0.01% of the soybean seed and 0.12% of the
jack bean seed dry weight was urease. Urease has been reported
to be present in non-leguminous seeds as well as in the seeds of
almost all species of legumes tested (7,150,180). The seed of
watermelon (Citrullus vulgaris) is reported to have 1/8th the
amount of urease found in jack bean seeds (199) or enough to
decompose its own weight in urea every hour (198). Urease also
occurs in various tissues of higher plants. Studies by
Granick in 1937 (72,73,74) on the distribution of urease in
seedlings of soybean and jack been demonstrated that the

cotyledons had the highest amount of urease, and that other

parts of the plant (roots, stem, and leaves) also contained
urease but at amounts less than one—tenth that found in the
cotyledons. Urease activity has been detected in the leaves of
the following vegetable plants: cucumber, bean, tomato, sweet
corn, celery and potato (88). Those plants which were most
easily injured by urea sprays, presumably due to the formation
of high concentrations of ammonium, had the highest urease
activity. Urease activities have also been detected in rice
plants (119), in the duckweed Spirodella (31). and in plant
tissue cultures (137,201).

Urease is also known to occur in two hundred species of
bacteria, in fungi, mollusks and crustaceans (183). The
production of urease by mammals is questionable (36), and its
occurrence in the gastric mucosa of several mammals is believed

to be bacterial in origin (189,38).

Physiological Significance of Urease in Plants

 

The most obvious function of urease is to hydrolyze urea.

A study which compared the pattern of incorporation of 14C into

amino acids in bean leaves fed 14C-urea or NaH14CO3 + NH4C1,

demonstrated that the pattern of incorporation was identical
for the two different nitrogen sources (188). These results
were consistent with the hypothesis that urea was metabolized by
its hydrolysis to 002 and ammonium. Thus when plants are given

urea as a nitrogen source, the enzyme urease plays an important

role in the assimilation of nitrogen.

10

Since urease is present in seeds in far greater quantities
than is needed for assimilation of exogenously applied urea,
alternative or additional functions for urease must be
considered. It has been suggested that the function of urease
in seeds is to serve as a storage protein (7,198). When the
amino acid composition of urease and reserve proteins were
compared, urease did not resemble the typical reserve protein
(7). Williams and Sharma (199) followed the changes in urease
and in total protein in the cotyledons of germinating CitruZZus
seeds. The urease activity was considerably higher on the 3rd
and 6th days of germination but dropped to nearly nondetectable
levels on the 9th day. During the decrease in urease levels,
there was no change in total protein content. The lack of
correlation between urease disappearance and breakdown of bulk
protein suggested that urease did not function as a reserve
protein. As an alternative to the storage protein hypothesis
these investigators proposed that the presence of urease in the
cotyledons was simply an indication of the state of the proto-
plasm of the cotyledons before their growth ceased in the
developing seed.

It is possible that the high urease content in seeds is
related to its role in the utilization of arginine and canavanine.
Arginine is a major amino acid in the storage proteins of cotton
(37), legume (97) and pine (202) seeds. In all three systems
the presence of arginine and its breakdown product urea was

correlated with the presence of arginase and urease activities

11

during the period of seed germination. Canavanine has been
detected in considerable amounts in the seeds of many economical-
ly important legumes (84) and serves as a principal nitrogen
storage metabolite for the developing plant (84,131,149,l79).
Canavanine utilization is initiated by an arginase-mediated
hydrolysis to canaline and urea (149,178), and urea is
subsequently converted to ammonium by urease. Rosenthal

studied the interrelationship of canavanine and urease in the
seeds of 29 species of canavanine-synthesizing legumes (150).

The seeds richest in urease generally had the most canavanine.
However, the urease content of the jack bean was disproportional-
1y greater than the quantity of stored canavanine. The massive
urease content of these seeds could not be rationalized as

being coupled to the size of the canavanine pool. Also, eight
species of Mucuna which contain urease in their seeds were
demonstrated not to contain canavanine as a storage compound.
However, an unknown amino acid which has properties similar to
canavanine was found in large quantities. Therefore the
suggestion that urease is required to hydrolyze the products of
canavanine breakdown is a valid assumption, but cannot be

accepted as the sole function.

Regulation of Urease

 

Since the urea concentration of plant cells is generally
very low (19), there may always be a small amount of urease

present to hydrolyze the urea formed from the degradation of

12

purines, arginine and other nitrogenous compounds. Most plant
tissue contains at least low levels of urease (72,73,74). When
urea is given as a source of nitrogen to the plant and the cells
are flooded with urea, higher amounts of urease would probably
be required. The question therefore arises: do urea and

the products of urea hydrolysis exert control over urease?

The earliest report of an increase in urease activity due
to application of urea in a plant was by Mokronosov et al. in
1965 (126). A 2 to 20 fold increase in the activity of urease
was observed in leaves of potato plants when leaves were treated
with urea solutions or when urea was introduced into the soil.

A year later, Matsumoto et al. (119) showed that rice plants
immersed in a urea solution doubled their urease activity in

2 hours and that inhibitors of RNA and protein synthesis
inhibited this rise in activity. However, the urease activity
decreased at 3 hours and plants grown on ammonium had activities
only slightly lower than plants grown on urea. They later
reported a similar rise and fall in urease activity with leaves
of jack bean, but this time the increase in activity was less
than double the original level (118). It was suggested that the
decrease in activity at 3 hours was due to a suppression of
urease synthesis by ammonium because addition of ammonium
resulted in a decrease in enzyme activity. Inhibitory action of
ammonium on urease in vitro was ruled out because ammonium was

removed from the extracts by dialysis.

13

Studies concerned with the metabolism of urea in Spirodela
oligorrhiza (duckweed) have contributed some interesting
information on the regulation of urease in this higher plant
(20,31). This organism is able to utilize urea as the sole
source of nitrogen only when the pH of the culture medium is
below 4.3. Growth on ammonium, nitrate or allantoin is not
similarly affected by pH. Urea uptake is not dependent on pH;
plants inoculated into urea medium at pH 6.4 exhibited no
growth and became nitrogen-deficient in appearance although they
contained about 100 ugrams of urea per gram fresh weight of
tissue. The detection of urease activity was correlated with
growth on urea. Plants with little or no urease activity soon
developed activities as high as 300 mUnits/gram fresh weight
after 20 days when placed in urea medium at pH 4.0. Urease
activities were even higher (500 mUnits/gr. fr. wt.) if the pH
was lowered to 3.5, even though 50 mUnits/gr. fr. wt. was an
adequate activity for normal growth on urea. Ammonium ions did
not inhibit urease activity and high levels of urease were
found in tissue containing a high ammonium concentration. When
allantoin was used as the nitrogen source, urease activities were
not affected by pH. Urease activity was also observed to rise
slowly when plants were left on nitrogen-deficient media; a
rise from 0 to 300 mUnits was observed during a 25 day period.
This was believed to be due to the formation of urea by the
breakdown of purines and arginine. It was evident that urease

activity appeared as a result of the presence of urea either from

14

an external or internal source. However, it was not resolved
why the use of exogenous urea was pH dependent but the use of
endogenous urea was not.

In a recent study by Polacco of urea assimilation in tissue
cultures of soybean (137), suspension cultures in the stationary
phase of growth supplied with a Murashige and Skoog (MS; ammonium
nitrate) nitrogen source were shown to have trace or zero levels
of urease. When these cells were transferred to fresh MS or
urea medium the urease levels increased, however the increase
was 10 to 20 times greater in the presence of urea. Ammonium
and methyl-ammonium inhibited the increase of urease activity in
the presence of urea by 33 and 60% respectively, but neither
compound inhibited soybean urease in vitro nor urea uptake.
Potassium citrate inhibited growth on urea but not on other
nitrogen sources, and completely inhibited the increase in
urease activity. Because urease from the jack bean has been
reported to contain 2 g-atoms of nickel per 105,000 g of enzyme
(39), it was suggested that the inhibition of the appearance of
urease by potassium citrate was due to its chelation of trace
Ni2+ in the growth medium.

In a later paper, Polacco (138) suggested the possible
requirement of nickel for urease because of the following

2

evidence. In the presence of urea, citrate and 10- mM NiSO4

‘growth was 800% greater than on urea alone. When the NiSO4

was replaced with CaCl2 or MgCl2 at concentrations of 5 or 10 mM,

growth was also greater than on urea alone, presumably because

15

these divalent cations were chelated by the citrate thus making
nickel available to the cells. At concentrations of NiSO4 higher
than 10.2 mM the citrate inhibition of growth on urea was not
alleviated. Citrate also inhibited the appearance of urease in
cell suspension cultures grown on arginine as the sole source
of nitrogen, and NiSO4 at 10"2 mM alleviated this inhibition.
Recently Polacco (139) has suggested that nickel is a
universal component of plant ureases because their levels were
stimulated when nickel was added to callus cultures of tobacco,
soybean and rice which were growing on solid MS medium containing
citrate. However, in this report citrate inhibited growth on
MS media of callus from all three species. This is contradictory
to the author's previous work (137,138) which documented that
citrate did not inhibit growth of soybean callus on MS media.
Also, growth of tobacco callus on urea was not inhibited by
citrate, suggesting that citrate inhibition of growth on urea
may occur only in soybean.

From the above studies it is evident that there is an
apparent induction of urease as a response to the presence of

urea. Ammonium may repress the synthesis of urease but has no

inhibitory effect on enzyme activity.

Extraction and Purification of Urease
Urease has a distinction in the field of protein chemistry
because it was the first enzume to be purified in crystalline

form (163). Sumner crystallized urease in 1926 by simply

16

extracting finely powdered, fat-free jack bean meal with 31.6%
acetone. The extract was filtered in an ice chest at 3 to 6°
and the filtrate was allowed to stand overnight. The crystals
formed were removed by centrifugation. Recrystallization
further purified the enzyme preparation (164). In 1941 Dounce
(42) established a more reliable recrystallization procedure
using an aqueous citrate-acetone extraction buffer. It was
difficult to assign a reliable specific activity because the
enzyme molecules had a tendency to aggregate and form oligomers
(71). Sumner's extraction procedure was eventually modified by
adding B-mercaptoethanol (144) to lessen aggregation, and EDTA
(17,151) to maintain a low concentration of metal ions. The
additions increased urease activity. Gel filtration on

Sephadex G-200, and ammonium sulfate precipitation also
increased specific activities (17). All of the above procedures
were developed for extraction of urease from jack bean meal.
Urease is commonly extracted from other plant tissues by
grinding the tissue in buffer plus a reducing agent or EDTA,
followed by centrifugation (20,119,137,201). The supernatant is

then assayed for urease.

Assays for Urease Activity

 

Techniques for the assay of urease activity have mainly
been based on the detection and measurement of the ammonium
released upon hydrolysis of urea. The simplest and oldest

assay employs an acid-base indicator which changes color in

17

response to the rise in pH caused by the presence of ammonium. A
more quantitative method is titration of the ammonium produced in
the enzyme reaction mixture (71,168,182). Blakeley et al. (17)
have proposed an elaborate assay procedure based on the same
principles in which a recording pH stat follows a linear
increase in pH with time. The most popular of the ammonium
detection methods is Nesslerization. The enzyme reaction is
stopped by addition of acid and the ammonium produced is
aerated off (182) or collected by absorption on a Dowex 50 resin
(103) and reacted with the Nessler's reagent which forms a
complex with ammonium that can be measured colorimetrically. The
ammonium must be removed from the reaction mixture because the
Nessler's reagent reacts with various other nitrogen compounds.
An assay which employs reactants that are considerably more
specific for ammonium nitrogen has been described by Kaplan (98).
Phenol and sodium hypochlorite in a basic medium, supplemented
with sodium nitroprusside as a catalyst, reacts with ammonium
to form an indophenol which absorbs light at 625 nm. Because of
its specificity for ammonium this assay can be performed
directly on the enzyme reaction mixture after the protein is
removed by precipitation (201). Urease activity has also been
determined by manometric measurement of carbon dioxide released
(181) and by measurement of 14CO2 released from 14C—urea (88).

A histochemical assay for urease was devised by Granick
(72). Haematoxylin, infused into plant tissue, changed color

from yellow to red in tissues where the pH was raised above

18

6.5 presumably due to the release of ammonium ion by urease. In
order to prevent ammonium from diffusing from the site of urease
action a complex was formed with the dye and a hydroxide of a
heavy metal which precipitated above pH 6.5. This precipitate
poisoned the enzyme which in turn stopped further liberation of
ammonium.

A staining procedure for detection of urease in gels has
been developed (54) which utilizes a tetrazolium salt that is
yellow, soluble and non-inhibitory to most enzymes in the
oxidized form, but upon reduction becomes blue and insoluble.
This dye will accept hydrogen from reducing agents above pH 8.
The urease is localized in the gel by the blue precipitate formed
when its alkaline product causes the tetrazolium to accept
hydrogen from a thiol. This staining procedure can only be used
on purified samples of urease because dehydrogenases will also

reduce the dye.

Catalytic Properties of Urease

 

The enzyme commission catalog (EC 3.5.1.5) lists the urease
reaction as urea + 2H20 = CO2 + 2NH3. From this equation it is
evident that two component reactions are involved resulting in
the breakage of two C-N bonds. Sumner et al. (169) have
demonstrated that when urea is hydrolyzed by crystalline urease
in the absence of buffer, ammonium carbonate is formed and then

is decomposed. This was illustrated by taking two aliquots from

the reaction solution and mixing with dilute alkali or with

19

dilute acid. The carbamate would remain in the alkali solution
but it would decompose to form ammonium carbonate in the acid.
The samples were then Nesslerized and the difference in the
colorimetric readings were equivalent to the carbamate
concentration. Sumner still did not rule out the possibility
that carbamate may be synthesized from carbon dioxide and ammonia.
More recently Gorin (69), carried out the urease reaction in the
presence of carbonic anhydrase, thereby establishing conditions
where recombination of ammonium and carbon dioxide could not
occur. Blakeley et al. (16) used a thymol blue buffer to give a
sensitive spectrophotometric measure of acidity changes. Both
reports have provided convincing evidence that carbamate is the

intermediate in a two-step reaction:
0 0

II +
+*H N-C-O-NH ] + HOH * H C0 + ZNH

u
urea + HOH + [H2N-C-OH + NH3 2 4 2 3 3

It is presumed that urease forms a carbamoyl complex as one of
the enzyme-substrate complexes and presumably water is the
acceptor in a carbamoyl transfer reaction, but no direct
evidence for this has been presented (16,142).

Sumner stated that urease is absolutely specific for urea
(166), but two other substrates have since been reported.
Urease catalyzes the hydrolysis of hydroxyurea, releasing
equimolar amounts of hydroxylamine, ammonium and carbon dioxide,
but the rate of hydrolysis decreases during the progress of
the reaction (59). Urease also catalyzes the degradation of

dihydroxyurea (55). Inhibition of hydrolysis of urea by

20

dihydroxyurea is less powerful than inhibition by hydroxyurea.
Reports on the effects of substrate concentration, pH and
temperature have been difficult to correlate and interpret
because of the discovery in 1951 that sodium and potassium ions
inhibit and phosphate ions activate urease (48). It has been
suggested that the real substances inhibiting urease are
complexes formed by the alkali cations with the phosphate
anions (103). It is not surprising then that the Michaelis
constants reported for the soybean and jack bean enzymes for
urea range from Km = 2.7 mM to Km.= 476 mM depending on the
buffer used (142). When TRIS sulfate buffer, which neither
inhibits nor activates urease, is used the Km is 4 mM (187).
The pH optimum has been reported to vary from 6.4 to 7.6 (166)
and in TRIS sulfate buffer it is 8 (187). The enzyme from
jack bean seeds is stable at temperatures as high as 65° (166).
Inhibitors of urease action include: reversible inhibitions
by thiourea (104); hydroxamic acid and its derivatives (16,106);
hydroxyurea (l6); and dihydroxyurea (55). Inhibition of urease
by its end product, ammonium, has been reported (90), however
others (137) have . demonstrated that it is not inhibitory.
Urease is inhibited by metal ions in the following order of
decreasing toxicity: Cu(II), Zn(II), Ni(II), Co(II), Fe(II) and
Mn(II) (156). The metal ions which form the most insoluble
sulfides are also the strongest inhibitors of urease (93,155),
which suggests that the inhibition is due to their reaction with

sulfhydryl groups near the active site (156).

21

Molecular Properties of Urease

 

The molecular weight of recrystallized urease, determined
by combining sedimentation velocity and diffusion data, was
reported by Sumner to be 483,000 D (167). A more recent study
reported the value, obtained by sedimentation equilibrium, to
be 489,000 D (143). Since most urease preparations are heter-
ogenous, these values pertain to the most prevalent molecular
species in freshly prepared urease. The molecular species
represented by this molecular weight is believed to contain 16
subunits (16n) of 30,000 D each (142), and it is possible to
observe the dissociation of the molecule in four discrete steps
to 8n, 4n, 2n and n. However, some controversy does exist as to
the subunit composition of the basic urease molecule. EM
studies by Fishbein have shown that the 8n species is a cyclic
trimer which combines with another trimer to form a spherical,
hexameric molecule of 480,000 D (56), thereby suggesting a sub—
unit number of six. The molecule represented by a MW of 260,000
D is enzymatically active and is formed by treatment with glycol
or glycerol (18,58) or by subjecting the enzyme solution to low
pH (70,171).

The 16n urease molecule has a propensity to aggregate,
forming larger molecular weight species. Studies of crystalline
urease using the ultracentrifuge demonstrated the presence of

three components possessing S values of 19, 28 and 36 S (34).

20,w

The distribution of protein among these components was not

affected by dilution, temperature change or standing, indicating

22

that the species involved were not in rapid equilibrium. In
the presence of sulfite, samples which previously gave three
ultracentrifugal peaks exhibited only the one at 19 8, suggesting
that the aggregates are held together by intermolecular disulfide
bonds. These aggregates, which are believed to be dimers and
trimers of the basic urease molecule and are enzymatically
active, have also been isolated by fractionation on CM-cellulose
(152), by elution from a Sephadex G-200 column (157) and by gel
electrophoresis (57,58). Aggregates representing molecular
weights as high as 6 times that of the 480,000 D species appear
as "strings of beads" under the electron microscope (56). No
physiological role has been suggested for these larger molecular
weight species of urease. Since all of the above mentioned
studies were conducted with urease extracted from seeds, these
polymers may represent a storage form of the enzyme.

The amino acid composition of urease has been reported
(125,143). However, these two citations are not in agreement.
Of particular interest are the values of 85 (143) and 66 (125)
half-cystines per 483,000 MW. Urease has a higher sulfur content

than seed storage proteins and average cellular proteins (7).

ATP-Dependent Hydrolysis of Urea -— The Allophanate Pathway

There are many species of green algae and yeasts which do
not contain urease but nevertheless can grow well on urea as a
sole source of nitrogen. How these organisms are able to

utilize urea had been a perplexing problem for many years. In

23

the early fifties, Walker (186) suggested that urea was utilized
via a reversal of the Krebs—Henseleit urea cycle because he
observed a stimulation of arginine formation in ChloreZZa
pyrenoidbsa grown on a urea medium. A similar suggestion was
proposed by Thomas and Krauss (175) because they too observed an
increase in arginine formation when Scenedesmus was given urea.
Hattori (77,78) obtained similar results with ChloreZZa
ellipsoidea. However, in a later paper (79), using 14C-urea,

he showed that 14CO2 was released and that there was incorpor—
ation of 14C into arginine. Hattori proposed a unified scheme
for urea and ammonium metabolism in this organism in which a
hypothetical compound A accepted ammonium or urea nitrogen with
formation of the compound A(NH2)2.

Kating (100) studied the metabolism of urea in two species
of yeast which lack urease, and proposed that urea is split into
a carbamyl and an amino group which are in turn metabolized by
transfer reactions to citrulline and glutamic acid. Cook and
Boulter (32) suggested a similar scheme for candida fiareri.
From the results of experiments in which 14C-urea was fed to
cells for short periods of time, they proposed that urea plus a
"C2" fragment gave rise to hydantoic acid which was then cleaved
to form carbamyl phosphate and glycine. Although all of the
above propositions were plausible to some extent, little
evidence was provided in their support.

In 1969, Hodson and Thompson (91) showed that Chlorella

vulgaris converted urea to CO and ammonium, but that ATP was

2

24

needed for the splitting of the urea molecule. At the same time,
Roon and Levenberg (146) described the presence of an ATP, Mg++
and K+ dependent hydrolysis of urea to CO2 and 2NH3 in extracts
of candida utilis, ChloreZZa eZZipsoidea and ChloreZZa

pyrenoidbsa. Potent inhibition of the reaction by avidin

suggested that the process was mediated by a biotin-containing

enzyme. The requirement of HCO3

(191) for this reaction proved
puzzling until it was shown that the hydrolysis of urea actually
involved two reactions: first, the carboxylation of urea by a
biotin-requiring enzyme in which a compound called allophanate
is formed; and second, the hydrolysis of allophanate to 2 C02
and 2 NH which requires neither ATP, Mg++, Rf nor biotin (147).

3
Thompson and Muenster (176) demonstrated that a crude extract of
ChloreZZa vulgaris possessing the above two reactions could be
separated physically by chromatography on brushite into two
enzymatic fractions: one which synthesized allophanate and
one which hydrolyzed allophanate. The two components involved
with urea hydrolysis in yeast are firmly bound together, resist
separation by a variety of methods and are believed to be a part
of a multienzyme complex (29,185,192,193). The reactions
involved can be distinguished from one another on the basis of
their sensitivities to heat, pH and chemdcal inhibitors. An

enzyme—CO complex, which is formed in the presence of all of

2
the cofactors required for carboxylation except urea, has been
isolated from saccharomyces cerevisiae by Sephadex chromatography

and is capable, in the absence of added cofactors, of

25

transferring the bound CO to urea (194).

2
A study which tested for urease or urea amidolyase (the urea

carboxylase-allophanate hydrolase enzyme system) activity in

extracts of unicellular algae representing five algal classes

showed that those algae classified as Chlorophyceae contained

urea amidolyase but no urease while all the other algae tested

contained urease but no urea amidolyase (111). The urea

amidolyase of candida utilis (148) and ChloreZZa (92) is

induced by urea, and the Km of the yeast enzyme for urea is

lO-fold lower than that reported for plant and bacterial

ureases. It has been suggested (148) that the lower Km for

urea might permit this organism to utilize urea at concentrations

which would be used very poorly by those species containing

urease. Attempts to find urea amidolyase in extracts from higher

plants have failed (148, 177).

Slow Changes in Phenotypic Expression

Organisms have the ability to make physiological changes to
accommodate for changes in their environment. Generally the
response begins immediately, particularly in the case of enzyme
formation where a certain enzyme is made in response to the
presence of its substrate. These types of changes can be turned
on or off depending on whether the stimulus is or is not there.
There are instances when the response is a gradual process which
eventually results in a metastable state which can gradually

revert back to the original state when the environmental stimulus

26

is taken away. This section will discuss situations where such
a slow change in phenotypic expression is known to occur. The
examples which follow all share certain characteristics: they
are changes which occur gradually over a long period of time;
once the change is expressed fully it will remain stable as
long as the environmental stimulus is present; and once the
stimulus is taken away the new expression is slowly lost.

An example of a gradual change in a phenotypic expression
in cultured plant cells is the change in the requirement for a
cell division factor in pith parenchyma from tobacco. These
cells require an exogenous source of a cytokinin for continuous
growth in culture on anMS medium (94) , but after a variable
number of transfers they sometimes lose this requirement (15,64).
After the requirement for cytokinin is lost the cultures are
said to be habituated and can be continuously cultured without
added cell division factors (15) which they are now able to
produce (45,46,200). This heritable change appears to be
epigenetic (132) rather than the result of a classical mutation
because it is directed, regularly reversible, leaves the cell
totipotent and involves the expression of a latent differentiated
function (15,64). A recent report by Meins and Binns (124)
used cloned cell lines derived from pith parenchyma of tobacco
to obtain evidence that this habituation process is gradual
rather than all-or-none and leads to progressively more
autotrophic tissues. Individual cells in culture showed

different degrees of habituation which were not phenotypically

27

fixed but could shift to higher or lower states of habituation.
These authors suggested that cytokinin habituation and autonomous
growth of crown gall tumor cells are analogous. Tumor formation
in crown gall involves the gradual and progressive change in
nutritional requirements for growth of the normal cell (23,24).
One event in this process is the conversion from requirement for
a cell division factor to a complete autotrOphy for this factor
which is a regularly reversible event (25). The events,
including cytokinin habituation, associated with tumor formation
are brought about by a self-replicating fragment of a plasmid
which is transferred from the crown-gall bacterium to the host
cell (205). Meins and Binns point out that cytokinin habituation
can occur in the absence of bacterial agents suggesting that this
aspect of tumor progression involves epigenetic cellular changes
of the type encountered in normal development. Other examples
(although not studied as completely as the above phenomenon) of
slowly acquired changes in cultures of plant tissue are: the
gradual development of salt tolerance in non-mutated cultures
of tobacco (129), the gradual loss of resistance to cycloheximide
in a cycloheximide resistant line of cultured tobacco cells (115)
and the loss of a requirement for auxin in cultured cells (64).
Studies in cultured animal cells have described a slow
adaptation phenomenon which involves an increase in the levels
of a specific enzyme. The resistance of various lines of animal
cell cultures to the 4—amino analogs of folic acid is often

associated with an increase in the cellular content of

28

dihydrofolate reductase (53,75,95,99,112,135). In murine
Sarcoma 180 cells, a sub-line (AT-3000) which produces larger
amounts of the enzyme was obtained by treating the cell
population with increasingly higher concentrations of methotrexate
(75). The response is gradual and variants are not obtained if
a high concentration of the drug is given at the onset of the
selection procedure. The dihydrofolate reductase present in the
AT-3000 cell line comprises as much as 6% of the total soluble
protein. This is an increase of ZOO-fold over the level in the
sensitive, parental cells (3). The higher amounts of the enzyme
are due to an increased rate of enzyme synthesis (3) which is
due to increased cellular levels of translatable dihydrofolate
reductase mRNA (101). When methotrexate resistant cells are
cultured in the absence of methotrexate the high levels of
resistance to the drug are lost (75). This loss in resistance
is correlated with a decrease in the rate of dihydrofolate
reductase synthesis (3) and the amount of the specific mRNA (lOlL
Instability is also a characteristic of methotrexate resistance
in other cell lines (14,95). Resistance and increased levels of
dihydrofolate reductase in certain lines of baby hamster kidney
cells (130) and murine leukemic cells (53) are stable when cells
are grown in the absence of the drug. A recent study by Alt

et a1. (2) reports the purification of cDNA sequences from the
AT-3000 cell line which are complementary to dihydrofolate
reductase mRNA. This cDNA was used to quantitate dihydrofolate

reductase gene copies in the variant cell line. Analysis of

29

the association kinetics of the purified cDNA with DNA from
sensitive and resistant cells indicated that the dihydrofolate
reductase gene was selectively multiplied approximately ZOO-fold
in the resistant line. Loss of resistance, when the cells were
cultured in the absence of methotrexate, was correlated with
loss of genes for the enzyme. After similar studies with stable
lines, it was concluded that selective multiplication of the
dihydrofolate reductase gene was responsible for the overproduc—
tion of dihydrofolate reductase in both stable and unstable
lines of methotrexate-resistant cells.. It was hypothesized (2)
that the gradual increase in the number of genes coding for
dihydrofolate reductase occurred because of unequal exchanges
between sister chromatids during crossingover.

The phenomena discussed abover are situations when
acquired traits have been passed on to subsequent generations.
There are two instances in higher plants when acquired traits
were reported to have been passed on to the offspring. The first
is the work of Durrant (43,44). Heritable changes were induced
in flax plants when they were supplied with different combina-
tions of nitrogen, phosphorus and potassium. The original
variety was changed into a larger form or a smaller form,
depending upon the fertilizer applied. The larger forms were 3
or more times heavier than the smaller varieties. The original
variety was intermediate in weight but taller than the others.
Both the large and the small forms were stable and remained

unchanged for seven generations, regardless of the fertilizers

30

subsequently supplied. The two new forms behaved as two

distinct genetic types when they were reciprocally grafted and
reciprocally crossed, giving no evidence of maternal or cyto-
plasmic inheritance. A similar environmentally induced heritable
change was reported by Highkin (85,86). It was found that
growth was inhibited in pea plants which were continuously
exposed to a constant temperature during the entire growth period
as opposed to normal conditions of fluctuating temperatures.

This inhibitory effect of constant temperature was cumulative
from generation to generation and reached saturation by the

fifth generation. The inhibition was reversible by returning
the plant line to fluctuating temperature conditions. However,
complete reversibility of the inhibition required at least

three generations of growth under the normal conditions.

MATERIALS AND METHODS

The Tobacco Cell Line and Growth Conditions

The XD line of cells used in these studies was originally
isolated from stem sections of Nicotiana tabacum L. cv. Xanthi
by Filner (50). The cells were grown at 28° C in 1 liter
flasks containing 500 ml of a chemically defined medium. The
flasks were shaken on a horizontal shaker which operated at 70
cycles per minute with a displacement of 3 inches. Subcultures
were routinely made by diluting aliquots of stationary phase
cultures (12 to 18 days old, depending upon the source of
nitrogen) 20-fold in fresh medium. The fresh weight of the

initial inoculum was 0.5-1.0 g/l.

Growth Media

 

Medium M-lD, which contains 2.5 mM nitrate, and nitrate-less
medium M-lD, in which equimolar amounts of the chlorides of
potassium and calcium were substituted for the corresponding
nitrates, were prepared as described by Filner (50,51). Ammonium
succinate M-lD was prepared by the addition of 3 mM ammonium
chloride and 1.5 mM succinic acid to nitrate-less M-lD. Casamino

acid M-lD was prepared by inclusion of 0.1% Casamino acids

31

32

(Difco Laboratories, Detroit, MI) to nitrate—less M—lD. Urea
M—lD was prepared by addition to nitrate-less M-lD of 0.3% v/v
of l M urea in water. All media were autoclaved for 20 min, and
the urea and Casamino acids stock solutions were sterilized by
filtration through a sterile membrane filter (Type HA, 0.22 um
pore, Millipore Corp., Bedford, MA). The urea was obtained from
two sources. Urea from Fisher Scientific Co. (Fair Lawn, NJ)
was first purified as described by Heimer and Filner (82) by
batchwise adsorption of anions on Bio-Rad AG 2—X8 ion exchange
resin in the hydroxyl form. Urea (ultra pure grade) obtained
from Schwarz/Mann (Orangeburge, NY) did not require purification
and was used directly. All media, and the urea and Casamino
acids stock solutions were adjusted with NaOH to pH 6.2 prior

to sterilization.

Harvesting of Cells and Preparation of Extracts

 

Cells were harvested by vacuum filtration on Whatman No. 1
filter paper and rinsed with deionized distilled water. After
determining the fresh weight, the cells were suspended in ice—
cold 50 mM bicine-NaOH (Sigma Chemical Co., St. Louis, M0)
buffer, pH 7.2, containing 5 mM dithioerythritol (DTE: Sigma).
Five ml of buffer were used per gram fresh weight. The cells
were homogenized by 30 strokes of a motor-driven teflon—glass
homogenizer (A. H. Thomas Co., Philadelphia, PA) at 4° C. The
homogenate was centrifuged at 10,000 rpm for 20 min at 4° C in a

Sorvall (Newton, CT) refrigerated centrifuge. The supernatant

33

was then centrifuged at 100,000 x g for l hr_at 4° C in a

Beckman (Fullerton, CA) ultracentrifuge using a type 65 rotor.
The resulting supernatant fraction was used as a crude enzyme
preparation after appropriate dilution with the homogenizing
buffer. Occasionally, this supernatant fraction was dialyzed in
2 l of 50 mM bicine-NaOH buffer (pH 7.2) with 0.5 mM DTE for

4-5 hr to remove ammonium ions. The procedure for preparation of
dialysis tubing (Scientific Products, McGraw Park, IL) was:

1) simmer in 50% ETOH for 1 hr, repeat; 2) simmer in 10 mM sodium
bicarbonate and lmM EDTA for 1 hr, repeat; 3) simmer in distilled
water for 1 hr, repeat. The dialysis tubing was stored in a

refrigerator in 50% glycerol.

Assay for Urease

 

Urease activity was determined by measuring the release of
ammonium ions when urea was incubated with the cell extract. The
complete reaction mixture contained: 50 mM bicine—NaOH (pH 7.2);
5 mM DTE; 50 mM urea; and from 1 to 600 mg protein of enzyme
extract. For each assay, 1 m1 of the reaction mixture was
prepared at ice temperature in a 10 ml test tube. The urea was
the last addition to the tube. The reaction was started by
placing the tubes in a water bath at 30° C. Incubation times
were 0 and 60 min. The reaction was terminated by adding 0.5 m1

of 5% w/v ZnSO followed by 0.5 ml of 0.2 M Ba(OH)2 (201). The

4

precipitate formed was sedimented by centrifugation at 1000 rpm

for 5 min and 1 ml of the resulting supernatant solution was

34

used for determination of ammonium ion concentration by the
phenol-hypochlorite method (procedure described below). Zero
time assays were used as controls. The increase in ammonium
ion concentration in the reaction mixture from 0 to 60 min was

taken as the measure of urease activity. Data are expressed in

+

umoles NH4

released per hour per gram fresh weight or per mg
protein. All assays were performed in duplicate. Reaction
mixtures which were heated in a boiling water bath for 5 min
prior to the 60 min incubation at 30° C exhibited no increase in
ammonium. The components of the reaction mixture, cell extract
and precipitating agents did not interfere with the ammonium
assay. When a cell extract was supplemented with a known
concentration of NH4C1 and was then subjected to the entire
assay procedure omitting the 60 min incubation, the value
obtained for the ammonium concentration (minus the amount
originally found in the extract) agreed to within :_S% with that

obtained with a standard of the same concentration dissolved in

water.

Assay for Ammonium Ion

 

Ammonium ion concentration was determined by a procedure
similar to that of Kaplan's (98). Two reagents were involved.
Reagent 1 consisted of 0.2 mM sodium nitroprusside in 1% w/v
phenol, and reagent 2 consisted of 0.125 N sodium hydroxide in
0.05% v/v sodium hypochlorite. The reaction was initiated by
adding successively to a l-md sample, 5 m1 of reagent 1 and 5 ml

of reagent 2. After being mixed on a vortex stirrer, the

35

solutions were incubated at room temperature for 30 min during
which time a blue color developed. Absorbance readings were

then taken at 625 nm on a Gilford (Oberlin, OH) spectrophotometer.
The absorbance reading of a blank which consisted of distilled
water plus the above reagents was subtracted from all values.

The concentration of the unknown sample was calculated from a
standard curve plotted from values obtained for known
concentrations of NH4C1. The reaction is consistent with Beer's
law for concentrations of ammonium ion from 0.01 to 0.50 pm;
samples with concentrations > 0.50 uM exhibit absorbance values

which increase linearly with increasing concentrations after

dilution with water.

Determination of Urea

 

The concentration of urea in cell extracts was determined

by first hydrolyzing it to NH;' and CO2 using a commercially
available urease and subsequently assaying for ammonium ions
using the procedure described above. Cell extract (0.5 ml) was
mixed with 0.5 m1 urease (1 mg/ml; purified from jack bean seeds;
specific activity 3200 units/gm, one unit will release 1 mg

NHJVS min at pH 7.0 and 30° C: Sigma Chemical Co.) and incubated
at 30° C for 60 min. The reaction was stopped with the same
precipitating agents used in the assay for urease. Ammonium ion
assays were performed on 0 and 60 min samples and the difference

between the two was used to determine urea concentration. The

urea concentration of unknown samples was calculated from a

36

standard curve plotted from values obtained for known

concentrations of urea hydrolyzed by urease.

Determination of Protein Content

 

Soluble protein content of the cell extract was determined
by the method of Lowry et al. (114). Protein in an 0.5 ml
aliquot of the cell extract was precipitated with 10% w/v
trichloroacetic acid, heated at 100° C for 5 min and sedimented.
The precipitate was washed twice with ice cold 95% ethanol, dried,
dissolved in l N NaOH and assayed for protein. Bovine serum

albumin dissolved in l N NaOH was used as a standard.

Cloning

The filaments of XD cells are uniseriate at the exponential
phase of growth (49). Each filament can be called a clone because
it is the result of a subdivision of a single cell. Therefore,
representative individuals from a cell population of a given
culture were cloned by isolating single filaments at the
exponential phase of growth. Each filament was grown on agar
medium until sizable callus tissue was formed and then
transferred to a liquid medium. Single filaments were isolated
by the following methods: a portion of the culture was poured
into a sterile petri dish and the smallest clumps visible with
the naked eye were collected one at a time with a 0.5 ml pipette
by capillary action; or single filaments were discerned with the

aid of a binocular dissecting microscope and collected one at a

37

time by pipetting. The first method resulted in the isolation
of filaments which were each composed of approximately 50 cells
or more. With the second method, filaments composed of 10 cells
or less could be easily isolated. The second method was used in
the later, critical work. However, it was discovered that
filaments consisting of 15 to 30 cells or more had the best
chance for survival (plating efficiency was sometimes as high as
80%) whereas filaments of less than 15 cells would not grow.
After transfer to agar medium, the clones took from 4 to 8 weeks
to grow to callus tissue with a diameter of approximately 10 mm.
The callus was then transferred into 5 to 40 ml of liquid medium
and allowed to grow under the culture conditions previously
described. A liquid suspension soon developed and the cultures
were maintained in the routine fashion. Urease assays were
performed on the cloned cultures no sooner than after the third

transfer on liquid medium.

RESULTS

Growth of Cells on Urea as the Only Source of Nitrogen

The ability of urea to support growth of XD cells was
examined by Heimer (80,81). He needed a reduced nitrogen
compound which would support growth but which was neutral in
the regulation of nitrate assimilation. Urea was suitable
because the cells grew when it was given as a sole source of
nitrogen, and they contained no detectable nitrate reductase.

The kinetics of cell growth on nitrate and urea are com-
pared in Figure l. The fresh weight and soluble protein
increased exponentially with culture age. The increase in
soluble protein was more rapid than the increase in fresh
weight for both sources of nitrogen. Growth on urea, in
comparison with growth on nitrate, was slower and yielded less
fresh weight at the end of the growth period. These cells were
previously grown on urea for at least 40 generations. The
doubling time for fresh weight increase was 2.5 days for the

cells grown on nitrate and 3 days for the cells grown on urea.

Activity of Urease in Cells Grown for many Generations on

Nitrate or Urea

 

Because the enzyme urease is assumed to be required for

growth on urea, cells grown on the two different nitrogen

38

39

Figure 1. Growth of XD cells on urea as the sole source

of nitrogen.

Stationary phase cells of a culture which had been
maintained on urea as the sole source of nitrogen for at
least 40 generations were subcultured into nitrate—less
M-lD medium supplemented with 3 mM urea. Fresh weight in
g/l (o) and soluble protein in mg/l (0) were determined
at various times during the culture period. The fresh
weight ( '0) and soluble protein (I) of stationary phase
XD cells maintained on M-lD and transferred‘to M-lD are

also included.

40

o 8.5 I 38:... SBEVEESn. oBBom

 

 

O
5
q

 

50'

IO"

o 3.5 a 295:3 \32935 amen.

 

20

I6

l2

Culture Age (Days)

41
sources were assayed for urease to determine if the presence of
urea in the medium had an effect on the amount of detectable

urease activity in the cells. The activities of urease measured

4.

in cultures grown on nitrate or urea, expressed in umoles NH4

formed per hour per gram fresh weight, are presented in Figure 2.
The cells grown on nitrate had specific activities of urease
ranging from 0.25 to 0.55 during the culture period, while
cells grown on urea (these cells were continuously maintained
on urea for 5 months prior to the experiment) had specific
activities ranging from 1.4 to 2.7. At all the times tested
during the culture period, cells grown on urea had higher
activities of urease than cells grown on nitrate. The highest
activity on both nitrogen sources was detected during the
exponential phase of growth. The urease activities did not
drop to zero during the stationary phase of growth. Cells
assayed between 0 and 4 days did not have activities of urease
higher than that found on the 4th day, i.e., there was not an
initial jump in urease activity at the time when cells were

transferred to fresh media. When the activities of urease are

+

4 formed/hour/

expressed in terms of soluble protein (Umoles NH
mg protein), the cells grown on urea still have the higher
activities (Figure 3). However, when the specific activities
are plotted in this manner urease is lowest during the
exponential phase of growth and highest at the stationary phase.
The rate of protein accumulation passes through a maximum in

the exponential phase of growth (49). Consequently, the

decrease in urease specific activity at this time is due to the

42

Figure 2. Specific activities of urease expressed on a
fresh weight basis for cells grown on urea or

nitrate.

Stationary phase cells which were previously maintained
on nitrate or urea for five months were subcultured into
fresh medium containing the same nitrogen source. The
activities of urease were assayed at various times during
the culture period for cells grown on urea (o) and nitrate

+. formed/

(D). The activities are expressed in umoles NH4

hr./gr. fr. wt.

43

 

 

l2

 

 

I.O*

- p n
5. o. 5.
2 2 I

:3 t “.9 .2 \ pogo... VIZ 3.9: 1 V 385

Culture Age( Days)

44

Figure 3. Specific activities of urease expressed on a
protein basis for cells grown on urea or

nitrate.

Stationary phase cells which were previously maintained
on nitrate or urea for five months were subcultured into
fresh medium containing the same nitrogen source. The
activities of urease were assayed at various times during
the culture period for cells grown on urea (o) and nitrate

+' formed/

(D). The activities are expressed in umoles NH4

hr./mg soluble protein.

45

 

 

 

. . _ _ P .
7. 6. 5. A 3 2 J
0 0 0 0 O O 0

2.295 95.2.3353 N12 «22:3 335

 

I2

Culture Age (Days)

46

higher production of proteins other than urease.

The apparent variation in urease activity with culture age
cannot be attributed to lack of reproducibility of cell
extraction. The reproducibility of the extraction procedure is
illustrated in Table I. One culture each of cells grown on
nitrate or urea was separated into three portions of 1 g each,
extracted and assayed for urease. The activities detected in
the 3 separate extracts from cells grown on each nitrogen source
did not vary appreciably.

A possible explanation for the lower urease activity in
cells grown on nitrate is that an inhibitor of urease may be
present in the extract of these cells. Table II shows the
results of 2 experiments where the extracts from cells grown
on urea and from cells grown on nitrate were mixed and assayed
for urease. In both experiments the activities of urease in the
two extracts were approximately additive, which rules out the
possibility of the presence of an activator or inhibitor in
either extract.

The rate of formation of ammonium catalyzed by urease was
constant with time (Figure 4A) and proportional to enzyme
concentration (Figure 4B) for extracts from both urea-grown and
nitrate-grown cells. The dependence of the rate of ammonium
formation upon urea concentration followed Michaelis-Menten
kinetics for enzyme from cells grown on either nitrogen source
(Figure 5A). When these data were plotted on a double

reciprocal plot (Lineweaver-Burk) an apparent Km for urea

47

Table I. Variability of activity of urease as measured with the

standard assaya.

 

Urease (Umoles NH; /hr./gr.fr.wt.)

 

Urea Nitrate
Extraction l 2.05 0.48
Extraction 2 2.10 0.47
Extraction 3 2.16 0.45

 

aOne culture each of cells grown on nitrate (5 days old) or
urea (15 days old) was separated into three portions of l g
each. These portions were then extracted and assayed for
urease according to the procedures described in the MATERIALS
AND METHODS section.

48

Table II. Effect of extract from cells grown on nitrate on

urease activity of extract from cells grown on

 

 

a
urea.
+
Urease (umoles NH4 /hr./gr. fr.wt.)
Nitrate Urea Mixed Calculated
Experiment 1 0.31 1.64 1.13 0.97
Experiment 2 0.54 2.12 1.38 1.33

 

aCells grown on nitrate or urea were extracted and assayed for
urease. The activity of urease was also determined for a
reaction mixture containing one—half volume each of the
nitrate and urea cell extracts. Included is a value s‘i;h
would be expected if one-half of the activity from each of the
nitrate and urea cell extracts was observed upon mixing.

49

Figure 4. Urease of cells grown on nitrate and urea:
proportionality with time and enzyme concen-

tration.

Cells in the exponential phase of growth on urea or
nitrate were extracted and assayed for urease. The
procedures were the same as described in the MATERIALS
AND METHODS section except that the time of incubation
and the amounts of extract added to the reaction mixture

were varied.

. . + . . .
A. Proportionality of NH4 formation with time of

incubation for extracts of cells grown on urea

(0) or nitrate (Cl).

. . + .
B. Proportionality of the rate of NH4 formation

with enzyme concentration for extracts of cells

grown on urea (o) or nitrate (t3).

50

 

 

 

 

 

 

g- o
'lu/peuuo; :HN sa|owrl

|.0

0.5
pl Cell Extract/lml Reaction Mixture

 

 

 

<

1
(D

J I

<1- N
m '1; 36/ peuua; :l-l N se|autrl

 

Time (hr)

51

. + .
Figure 5. Dependence of the rate of NH4 formation on urea
concentration in extracts of cells grown on urea

or nitrate.

Cells in the exponential phase of growth on urea or
nitrate were extracted and assayed for urease. The complete
assay mixture was prepared as described in MATERIALS AND
METHODS with various concentrations of urea.

+
4
concentration for extracts of cells grown on

A. Dependence of the rate of NH formation on urea

urea (0) or nitrate (:3).

B. Double reciprocal plot (Lineweaver—Burk plot) of
+ .
the dependence of the rate of NH4 formation on
urea concentration; urea cell extract (a) and

nitrate cell extract ((3).

52

 

 

 

|.0

 

 

655*

Urea (mM)

0.l

 

 

71.2312 «22...:

 

 

20

IO
mM Urea"|

 

 

p
0
2

p
O

Tofuxz 3.05.:

-Km '

53

of 0.20 mM was calculated for the enzyme obtained from both
types of cells (Figure 5B). This value is lower than the Km
measured for urease from other plant sources (142). The above
results are consistent with the idea that the urease molecules
extracted from cells grown on the two different nitrogen
sources are the same, but are present in different amounts.

The difference in the activities of urease between cells
grown on nitrate or urea were consistently observed. Figure 6
depicts the urease activity in the two types of cells over a
time span of approximately one year. The transfer periods were
12 to 14 days for the cultures grown on nitrate and 16 to 18
days for the cultures grown on urea. Each point on the graph
represents either an activity determined during the exponential
phase of growth or an average activity calculated from values
obtained at different ages of the culture. During the year,
the urease levels of the cells grown on nitrate varied little
from the specific activity of 0.5. The urease levels varied
considerably in the cells grown on urea during this period, but
they never dropped to levels which were equal to those found in

the cultures grown on nitrate.

Urease Activity of Cells Transferred from Nitrate Medium to

Urea Medium

 

Since it has been reported that urease activity increases
in plant tissue when urea is given as a nitrogen source (20,119,

126,137), it was of interest to determine if the higher levels

54

Figure 6. Differences in the activities of urease in
cultures grown on nitrate or urea over the

period of a year.

Cultures maintained on urea (0) or nitrate (:3) were
assayed for urease at various transfers over the period of
a year. Each point on the graph represents either an
activity determined during the exponential phase of growth
or an average activity calculated from values obtained
at different ages of the culture. The transfer periods
represent 12 to 14 days for the cultures grown on nitrate

and 16 to 18 days for the cultures grown on urea.

55

 

 

 

2‘0

 

 

 

2.0l' °

0

'0.
0
(1m; 36/ au/ pauuo; :HN salowrl ) asaein

I5

IO

Transfer

56

of urease detected in the cultures grown on urea were due to

an induction of urease synthesis. Cells, previously grown on
nitrate and in the stationary phase of growth, were transferred
to either nitrate or urea medium and were assayed for urease

at various times during the culture period (Figure 7). The
activities of urease for the cells grown on urea were similar
to those of the cells grown on nitrate during the first 4 days
of the culture period. After this time the activities of the
urea cells were higher than thosecfifthe nitrate cells. However,
the specific activities of the cells on urea never exceeded

0.5, i.e., the activities of urease were never higher than

what is normally found in cells grown on nitrate. When
directly transferred from nitrate medium the cells grew on urea,
but the rate of growth and the yield in fresh weight at the end
of the growth period were approximately one-half the normal
values for cells grown on nitrate. The slower growth rate and
consistently lower levels of urease in cells newly transferred
from nitrate to urea was not due to a lack of urea uptake.
Figure 8 compares the cellular urea concentration, at various
times during the age of the cultures, of cells newly transferred
from nitrate to urea medium and of cells previously grown on
urea for more than a year. The urea concentration was higher
in the cells which were previously cultured on nitrate. Cells
which were newly transferred into urea medium had the ability
to accumulate urea, and the higher amounts of urea in these

cells compared to urea-conditioned cells may indicate that the

57

Figure 7. The levels of urease in cells transferred from

nitrate to urea medium.

Cells in the stationary phase of growth on nitrate
were transferred to a medium containing urea or nitrate
as the sole source of nitrogen. At various times during
the culture period the cells growing on urea (o) and

nitrate (:3) were assayed for urease.

58

 

 

 

_ _ b _ _

 

 

 

5. 0. 5. 0. 5.
2 2 I I 0
:seteefomgoa ”:2 3.9: 1:38:

l4

IO

Culture Age (days)

59

Figure 8. Urea concentration of cells consistently
maintained on urea and of cells newly transferred

into urea medium.

Cells in the stationary phase of growth were
transferred to a medium containing urea as the sole source
of nitrogen. At various times during the culture period
the cells were extracted and the urea concentration of the
cell extract was determined as described in MATERIALS AND
METHODS. The urea concentration is expressed in umoles/
gr.fr.wt. for cells previously maintained on urea for many
generations (0) and for cells newly transferred from

nitrate medium into urea medium (:3).

60

 

 

 

1 I l I

0. 00. m. <1: N
— o o o o

( 'lM '1; ‘10/ salow 1'!) ann

IO

In I

Culture Age (days)

 

61

lower levels of urease are insufficient for adequate hydrolysis
of cellular urea.

When cells previously grown on nitrate were consistently
cultured on urea medium their growth rate increased on subsequent
culturings up to a doubling time of approximately 4 days, but
their levels of urease remained low for many subcultures. When
these cells were continuously maintained on urea medium the
activities of urease eventually increased. Figure 9 compares
the activity of urease at the exponential phase of each transfer
period for cells transferred from nitrate to urea and for cells
maintained on nitrate. The urease levels remained the same for
both types of cells for the first three transfer periods, but
at the 8th transfer the cells on urea had an activity of urease
which was 5 times higher than that of the cells on nitrate. A
decrease in the urea concentration of these cells was correlated
with an increase in urease levels. The urea concentration of
the cells of all the cultures grown on urea which exhibited
the lower levels of urease in this long term experiment were
5 times higher than the urea concentrations of the cells
possessing the higher levels of urease. When this long term
experiment was repeated (Figure 10) there was a similar
increase in the levels of urease in the cells grown on urea.

In the second experiment there was a gradual increase in the
specific activity of 2.0 at the 15th transfer. Although in

both experiments the urease levels increased, the time taken‘

62

Figure 9. Long Term Experiment No. l: urease activity of
cells transferred from nitrate to urea medium

and consistently maintained on urea thereafter.

Cells previously grown on nitrate and in the
stationary phase of growth were transferred into urea or
nitrate medium. These cultures were continually maintained
on the two different media for 11 transfer periods. At
the exponential phase of growth of the first 3 and last 4
transfer periods the cells were assayed for urease.

+formed/hr./gr.fr.wt. for

4
the cells maintained on urea (o) and nitrate (:3).

Values present are in umoles NH

63

 

\D—"‘

ll—

/i/--°\

 

 

 

h h b h
5 0 5. 0 5
2 2 I I O

:3 he .19 “.5885. .312 8.683 0883

l0 ll

8 9

3
Transfer

2

64

Figure 10. Long Term Experiment No. 2: urease activity
of cells transferred from nitrate to urea
medium and consistently maintained on urea

thereafter.

Cells previously grown on nitrate were transferred in
the stationary phase of growth into urea or nitrate medium.
These cultures were continuously maintained on the two
different media for 18 transfer periods. The cells were
assayed for urease during the exponential phase of growth.
Values presented are in umoles NH; formed/hr./gr.fr.wt.

for the cells maintained on urea (o) and nitrate (c3).

65

 

 

 

 

 

0. In.
N .—

(‘lM'Jl'be'Ju/pemio;:HN saloer) asaain

LO-
.5

I5

[0
Transfer

66

to reach the higher levels normally found in urea-conditioned
cells was different. In the first experiment it took at least
120 days before the cells exhibited the high urease activities,
while in the second experiment the increase occurred in 220 days.
In both experiments, the high levels of urease were consistently

maintained after the initial increase.

Is the Increase in Urease Due to the Selection of a Spontaneous

 

Variant?

The slow increase in urease activity in cells which were
transferred from nitrate to urea medium could be due to the
selection of a variant cell which has a high urease activity.
This does not appear to be true for the following reasons.

A selection of a variant able to grow among a population of
cells unable to grow on urea is not occurring here. The
growth rate is slower than that on nitrate, but a dramatic
death of the culture followed by a slow appearance of growth
after 2 or 3 months is not observed. Such massive culture
death and subsequent growth has been observed when XD cells
were selected for resistance to threonine (80,81) and to lysine,
methionine and proline analogs (196). The frequency of
spontaneous variants (presumably mutants) has been estimated
to be about 10'.7 in a normal population of these cells, based
on the fact that about ten variant cultures grow up when a
hundred cultures containing 106 cells each are subjected to a

strong selection (195, P. Filner personal communication).

67

Because the rate of growth on urea is slightly faster for
high-urease cells than for low-urease cells (see section on
comparison of growth rates), it would take a spontaneous
variant which has the high levels of urease 500 days to take
over a low urease producing cell population. Since the actual
observed times for the increase in urease was 120 and 220 days,
it is unlikely that the increase was due to a selection of a

variant cell.

Is the Increase in Urease Due to the Selection of a Large

 

Pre-existing Subpopulation of High-Urease Cells?

 

The slow increase in urease activity in cells transferred
from nitrate to urea could be due to a gradual selection of a
pre-existing subpopulation of high urease producing cells
during the many transfer periods, provided that these cells
are sufficiently abundant. In order to determine if such high-
urease cells exist in a population of cells grown on nitrate,
clones were isolated from this population and separately assayed
for urease. The solid lined histogram in Figure 11 represents
the urease activities of 83 such clones. The urease activities
of these clones were rather narrowly distributed around the
average value normally found in the uncloned population of
nitrate-grown cells (solid lined arrow). The highest specific
activity observed was 0.8. None of the clones has an activity
near the average value present in urea-conditioned cultures

(dashed arrow). The average specific activity of these clones

68

Figure 11. Urease activities of clones derived from
populations of cells having low and high

levels of urease.

Clones were derived by culturing single filaments on
solid medium. After the filaments grew to sizable callus
tissue they were transferred to liquid media and maintained
in liquid culture. The clones were assayed for urease
during the exponential phase of growth. The solid lined
histogram represents the urease activity of 83 clones
derived from a population of cells grown on nitrate as the
sole source of nitrogen. The dash lined histogram
represents the urease activity of 26 clones derived from a
population of cells previously grown on urea for many
generations. The solid arrow and dashed arrow indicate
the average values for urease activity normally found in
the entire population of cells grown continuously on

nitrate and urea, respectively.

69

A..3.t.to\...;\ 665.3,. H12 3.25.: 335

 

 

. . .2

'0“ .1000‘ n c Jo ”

r I. c n o
p-.. r-.. rL

.

o

.

o

o

n

o

-

. . . . O.—
.r ..... . I. -.. . C _I

 

 

 

 

 

 

 

seuolg 40 JaqumN

70

of nitrate-grown cells was 0.34 and the standard deviation was
0.154 (1'45%). Since the variation in activities appears to
follow a bell—shaped curve, one can use a table of areas of the
normal curve (5) which is derived from the integral of the

probability function

 

x
1 I -t2/2
[P(x) = V2fl ie dt, where x is the standard deviatioé]

to calculate what percent of the cell population has urease
activities above or below a certain level. For example, the
percent of the cell population which is predicted to have urease
specific activities between 0.34 and 0.80 is 49.8%. From this
percentage we can calculate that for the entire population, 1
in 200 cells would be expected to have an activity greater than
0.8. Therefore, the fact that one clone was found with an
activity of 0.8 does not seem unreasonable. In a similar
manner, it was calculated that the probability of a cell

having a specific activity greater than 1.0 is l in 33,000.
Determination of the fraction of the population having a urease
activity greater than 2.0 is more difficult because this is

10 standard deviations from the mean. Regular tables give only
areas for the normal curve for values which are less than or
equal to 4 standard deviations from the mean. However, a table
is available for large values of x which was computed using an
asymptotic expansion of the probability function (113). From
this table it was calculated that approximately 1 in l x 1023

cells could have an activity of urease greater than 2.0.

71

Since cultures at stationary phase contain 108 cells per liter,
one can conclude that it is highly unlikely that even a single
individual in the population of cells grown on nitrate has a
high activity of urease. Using the difference in the rates of
growth for high and low urease cells (see section on comparison
of growth rates) and the observed times in which the increase
in urease occurred, the initial number of high urease cells
required to be present in the cell population was calculated
to be 1 cell in 9 in order for the increase to occur in 120 days
and 1 cell in 500 in order for the increase to occur in 220
days. Clearly the increase in urease cannot be attributed to
a selection of a pre-existing subpopulation of high-urease
cells.

Clones isolated from a urea-conditioned cell population
also exhibited varied urease activities. The isolation of a
large number of clones from a urea-conditioned cell population
was difficult because the plating efficiency was less than 1%
for filaments cultured on urea supplemented agar medium. The
plating efficiency was slightly improved when urea-conditioned
filaments were cultured on nitrate agar medium. Therefore,
most of the clones isolated from the urea-conditioned cell
population were first grown on nitrate agar for 6-8 weeks and
then transferred to liquid urea medium. The dashed lined
histogram in Figure 11 represents the activities of urease
found in 26 clones —- 5 initially grown on urea agar and 21

initially grown on nitrate agar -— derived from a urea-conditioned

72

cell population which produced the higher levels of urease.

The specific activities of the clones varied from the average
specific activity normally found in urea-conditioned cells

(the dashed arrow), but activities similar to those found in
nitrate-conditioned cells were not observed. The lowest
specific activity detected in a clone was 1.60. The average
specific activity of these clones of urea-conditioned cells was
2.43 and the standard deviation was 0.647 (:_26%). Some clones
had specific activities which were nearly double that found, on
the average, in urea-conditioned cells. The distribution of
urease activity in the clones initially cultured on urea agar
was similar to the distribution in the clones initially cultured

on nitrate agar.

Increase in Urease Activity Durinngrowth on Urea in Clones

 

Derived from a Population of Cells Previously Grown on Nitrate

 

If the increase in urease is due to the selection of a
single spontaneous variant or a pre—existing subpopulation of
high urease producing cells, one would expect that some
nitrate-conditioned clones would exhibit an increasethurease
when transferred to urea medium and some would not. Eight
clones were isolated from a population of cells grown on
nitrate, cultured on urea agar, and subsequently maintained
in liquid medium containing urea as the nitrogen source. At
the exponential phase of growth of the 5th and 10th transfers

they were assayed for urease (Table III). The urease activity

73

Table III. Urease activity of clones isolated from a
population of cells grown on nitrate and transferred

. a
to urea medium.

 

Urease (umoles NHZ?hr./gr.fr.wt.) at:

Clone Number 5th Transfer 10th Transfer

 

1 0.38 0.73
2 0.68 0.96
3 0.48 0.97
4 0.43 0.99
5 0.59 0.92
6 0.55 1.48
7 0.67 0.87
8 0.51 0.89

 

“Single filaments were isolated from a culture in the exponen-
tial phase (4 days old) of growth on nitrate and grown on agar
medium containing urea as the sole source of nitrogen. After
7 weeks, the callus tissue formed was cultured in liquid
medium containing urea. The clones were assayed for urease at
the exponential phase of growth after 10 and 20 weeks (5th and
10th transfer periods) of growth on liquid medium. The
urease activity of the original uncloned parent culture was
0.45. The average urease activity at the 5th transfer was
0.54 with a standard deviation of 0.14 (:_26%). The average
urease activity at the 10th transfer was 0.98 with a standard
deviation of 0.22 (i_22%).

74

increased in all of the clones from the 5th to the 10th transfer.
In 4 of the clones, the urease activity at the 10th transfer was
double the activity at the 5th transfer. It appeared that all of
the clones were in the process of adapting to the higher levels
of urease. Since all clones exhibit roughly comparable

increases in levels of urease, the hypothetical selections

described above are not occurring.

Comparison of Growth Rates of Cultures which Exhibit the High
or Low Levels of Urease and Calculations for Rise in Urease

Due to Growth of High—Urease Cells

 

Urea-conditioned cells which have the higher level of
urease grow faster on urea than cells with the lower levels of
urease. Figure 12 compares the difference in growth of the
following types of cells: cells which were grown on urea for
many generations and have the high levels of urease: cells
which were transferred from nitrate medium to urea medium,
grown on urea for three transfer periods and still have the
low levels of urease: and cells which were transferred from
nitrate to urea medium for the first time. The growth rate and
doubling time for the entire culture period of these cells

can be calculated using the following equations:

 

 

t
1n
C
o lni?
K = --—-- and T =
t d K

where K = rate constant for growth, Ct = fresh weight at day15,

75

Figure 12. Differences in growth on urea of high-urease

and low-urease cells.

Cells in the stationary phase of growth were weighed
under sterile conditions so that all inocula were identical
and transferred to fresh medium containing urea as the sole
source of nitrogen. At 3 day intervals during the growth
period triplicate samples of the cultures were harvested
and the fresh weight determined. Fresh weight is plotted
on a log scale versus culture age for: urea conditioned
cells which possess the higher levels of urease (0);
cells recently transferred from nitrate medium to urea
medium and maintained on urea for 3 transfers, which
possess the lower levels of urease (c3); and cells newly
transferred from nitrate to urea which possess the lower

levels of urease (A).

76

 

Fresh Weight (g/L)
5

 

l l I I

I.

 

 

3 6 9 I2
Culture Age (Days)

I5

77

C
0

initial fresh weight of cells, t = time (15 days), and
Td = doubling time. The growth rate constants and doubling times
for the three types of cells were 0.230 days-1 and 3.0 days for
the high-urease cells, 0.193 days.-1 and 3.6 days for the low-
urease cells, and 0.136 days-1 and 5.1 days for the cells newly
transferred from nitrate to urea. The difference in the rates
of growth of the high-urease and low—urease cells on urea were
consistently observed for the cultures used in all of the
experiments in this thesis.

Because the growth of cells which have the higher levels of
urease is faster than cells with the lower levels of urease,
one can use the growth rate constants which were calculated
above to determine how long it would take high-urease cells to
take over and dominate a low-urease cell population grown on
urea. It was shown earlier in this thesis (p. 70) that 1 cell
in 33,000 in a population of cells grown on nitrate would be
expected to have a urease specific activity greater than 1.0.
Using the growth rate constants for the high—urease and low—
urease cells, it was calculated that it would take 350 days
for 93% of the cell population to be composed of cells with an
activity greater than 1.0. Since cells transferred from nitrate
to urea reached twice this value of urease specific activity in
120 and 220 days (Figures 9 and 10) it is unlikely that the
increase in urease was due to the gradual takeover of high-
urease cells in the population, provided that the rate of

growth and the increase in urease is not affected in mixed

78

populations of high and low urease cells.

Effect of Mixing Nitrate-Conditioned and Urea-Conditioned Cells

 

on the Rise in Urease Activity

 

The following experiment was performed to determine if
low-urease cells and high-urease cells behave independently in
mixed populations and if the high-urease cells have an effect
on the increase in urease of lowburease cells. Stationary phase
cultures of urea—conditioned and nitrate-conditioned cells were
transferred to urea medium. Mixtures of these cells at
approximate ratios of 50 urea to 50 nitrate and 10 urea to 90
nitrate were also transferred to urea medium. These cultures
were maintained for three transfer periods and assayed for
urease at the exponential phase of growth for each period. The
results of this experiment are presented in Table IV. Included
in this table are the estimated ratios of the make—up of the
cell population in the mixed cultures at the time when urease
was assayed, i.e., the percent nitrate and urea cells present.
Since the two cell types grow at different rates, the ratios
should change. The estimated ratios were calculated by using
the growth rate constants for the three different cell types
reported in the previous section. The growth rate constant of
0.230 days.-1 was used for the urea cells whereas the growth
rate constant of 0.136 days71 was used for the nitrate cells of
the first transfer period and 0.193 days.-1 was used for the

nitrate cells of the 2nd and 3rd transfer periods. Also

79

Table IV. Urease activity of mixtures of urea-conditioned cells

and nitrate-conditioned cells cultured on urea

 

 

mediumq
b New Calculated
f
Trans er Cells Urease Ratioc Ureased
lst Urea 2.66
50 U:50 N 1.64 65 0:35 N 1.90
10 U:90 N 0.89 14 U:86 N 0.79
Nitrate 0.49
2nd Urea 2.50
50 U:50 N 1.90 87 U:13 N 2.25
10 U:90 N 1.35 42 0:68 N 1.47
Nitrate 0.62
3rd Urea 2.46
50 U:50 N 2.00 92 U:8 N 2.22
10 U:90 N 1.90 55 U:45 N 1.60
Nitrate 0.59 I

 

aStationary phase cells previously grown on nitrate (N) or urea
(U) were cultured into urea medium at ratios of l U:0 N, 50 U:
50 N, 10 U:90 N, and 0 U:l N. At stationary phase the cultures
were transferred to fresh urea medium for 3 transfer periods.
The cultures were assayed for urease during the exponential
phase of growth at each transfer period.

g'formed/hr./gr.fr.wt.

cSince the rates of growth on urea are different for the nitrate
and urea conditioned cells, the change in the ratio of nitrate
to urea cells brought about by the difference in the growth
rates was calculated for the day of assay using the growth rate
constants determined from Figure 12.

d

An expected activity was calculated from the activity observed
in the nitrate and urea cell cultures and the ratio of urea
cells:nitrate cells determined for the mixed cultures at that
transfer period.

umoles NH

80

included in the table are the predicted values for activities of
urease which would be present if the cultures did contain the
calculated ratios of nitrate:urea cells. In all cases, the
calculated activities were not appreciably different from the
observed activities of the mixed cultures. The presence of
neither 10% nor 50% high-urease cells caused the low-urease
cells to develop high levels of urease. These results are
consistent with the interpretation that the activities of high—
urease and low-urease cells are independent of one another and
that these cells grow independently of one another in mixed

populations.

The Effect of Various Nitrogen Sources on the Activity of Urease

 

in Urea-Conditioned Cells

 

Another test of the possibility that the high-urease cells
result from a selection process is to determine stability of the
phenotypes in the absence of the putative selection agent. The
stability of the high urease levels in urea-conditioned cells
was tested by transferring these cells to media which contained
alternate sources of nitrogen. When urea-conditioned cells were
transferred to a medium which contained nitrate as the sole
source of nitrogen the activities of urease were lower in
these cells than in similar cells which were recultured on
urea (Figure 13). When urea was given in conjunction with
nitrate, the activity of urease decreased in a similar manner.

The levels of urease in the cells cultured on nitrate were

81

Figure 13. Decrease in urease activity in ureawconditioned

cells transferred to nitrate medium.

Urea-conditioned cells in the stationary phase of
growth were transferred to urea or nitrate medium. The
cells were assayed for urease at various times during the

culture period. Urease activity is expressed in umoles

+

NH4 formed/hr./gr.fr.wt. for the cells grown on urea (o)

and nitrate (t3).

82

 

 

l J
8 I2

4
Culture Age (days)

 

- h - p P

 

-
5. o. 5. o. 5.
2 2 I I 0

63.2%..325 3E3.— HIz 3.9: 1:82:

83

higher than the levels normally found in nitrate conditioned
cells. The cells were making considerably high levels of urease
even though urea was not present in the culture medium. These
cells were continuously cultured on nitrate medium and assayed
for urease at the exponential phase of growth of each transfer
period (Figure 14). The urease activity gradually decreased in
these cells during the first four transfers until a specific
activity of 1.0 was obtained. After the 4th transfer these
cells appeared to have a urease specific activity which had
stabilized around 1.0. The gradual and parallel rises observed
after the 11th transfer in both cultures probably have nothing
to do with the nitrogen source. For unknown reasons, about that
time cultures began growing to slightly higher yields suggesting
a change in some uncontrollable environmental parameter, e.g.,
traces of growth inhibitors may not have been present in new
chemicals used to make media. This is one of the unfortunate
hazards associated with experiments done over a long time span.
The lower activity in the cells grown on nitrate could not be
due to an in vitro inhibition of urease by nitrate present in
the cell extract because a previous experiment had shown that
mixing of nitrate and urea cell extracts had no effect on the
activity of urease (Table II). Nitrate exerts an effect on
the activity of urease in urea-conditioned cells by causing a
gradual decrease in activity. However, the activities only
decrease to levels which are twice that found in nitrate-

conditioned cells.

84

Figure 14. Gradual decrease in urease activity in urea—
conditioned cells transferred to nitrate

medium and maintained on nitrate medium.

Urea-conditioned cells in the stationary phase of
growth were transferred to urea or nitrate medium. The
cultures were maintained on the same nitrogen source for
19 transfer periods. The cultures were assayed for urease
at the exponential phase of growth for most of the
transfer periods. Urease activity is expressed in umoles
NH+ formed/hr./gr.fr.wt. for the cells grown on urea (o)

4
and nitrate (E1).

85

 

 

 

-

 

 

 

- n
0 5. O 5.
I I 0

23 E u0\.a.._\uoEt£ lwIz 8.083 3095

20

IO
Transfer

86

A similar experiment was performed using ammonium succinate
as the nitrogen source. During the first transfer period
(Figure 15), the cells grown on ammonium succinate had activities
which were less than or equal to one-half that of the activity
found in similar cells recultured in urea. When cells were
cultured in a medium containing both urea and ammonium succinate
(3 mM each) the activities decreased in a similar manner. When
urea-conditioned cells were continuously cultured on ammonium
succinate (Figure 16) the specific activity of urease at the
exponential phase of growth eventually decreased to 0.5 on the
9th transfer. The cells continued to exhibit this low level
of urease activity for the next 7 transfers. The ammonium
present in extracts from cells grown on ammonium succinate did
not appear to have an effect on the activity of urease for the
following reasons: 1) removal of ammonium succinate from
the extracts of ammonium and urea grown cells by dialysis did
not result in an increase in activity of urease; 2) the
ammonium concentrations of extracts from the cells grown on
urea (determined from the zero time point of the urease assay)
were equal to or slightly higher than the ammonium concentra-
tions of extracts from the cells grown on ammonium succinate:
and 3) the urea concentration used for the urease assay was
approximately lOO-fold higher than the ammonium concentration
in the cell extracts. When the ammonium succinate grown cells,
which were now making the low levels of urease, were transferred

back to urea medium it took three transfer periods before they

87

Figure 15. Decrease in urease activity in urea-conditioned

cells transferred to ammonium succinate medium.

Urea-conditioned cells in the stationary phase of
growth were transferred to urea or ammonium succinate
medium. The cells were assayed for urease at various

times during the culture period. Urease activity is
+
4
cells grown on urea (o) and ammonium succinate (:3).

expressed in umoles NH formed/hr./gr.fr.wt. for the

88

 

 

 

 

 

 

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1.0..
\m
y
a
d
e
9
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r
U
H...
U
C
v

_ F a _

5. 0. 5. O 5

2 2 I I. 0.

A .§.¢...o\.2\ooE.£ HIZ 3.2: .3 $35

89

Figure 16. Gradual decrease in urease activity in urea—
conditioned cells transferred to ammonium
succinate medium and maintained on ammonium

succinate.

Urea-conditioned cells in the stationary phase of
growth were transferred to urea or ammonium succinate
medium. The cultures were maintained on the same nitrogen
source for 16 transfer periods. The cultures were assayed
for urease at the exponential phase of growth for most of

the transfer periods. Urease activity is expressed in
+

4
urea (o) and ammonium succinate (El).

umoles NH formed/hr./gr.fr.wt. for the cells grown on

 

90

 

 

O
" O
O
O
J
0. '0.
IO N

I0

 

 

 

('Imnb/uu/peuuo; :l-IN se|omrI ) esaein

Transfer

91

were again making the higher levels of urease (Figure 17). This
increase in urease was considerably faster than the increase
detected in cells which were transferred from nitrate to urea,
that had never been exposed to urea previously. The cells

grown on ammonium succinate "remembered" that they had previously
grown on urea. It would be interesting to test if cells
transferred from nitrate to ammonium succinate, grown on
ammonium succinate for an extended period of time, and then
transferred to urea would behave as the ammonium succinate cells
in this experiment (rise in urease in 3 transfers) or behave as
nitrate cells which had never been exposed to urea (rise in
urease only after many transfers). Unfortunately this
experiment was not conducted.

Casamino acids will also cause the urease levels in urea-
conditioned cells to decrease. Figure 18 gives the activities
of urease for urea-conditioned cells cultured on urea or
casamino acids medium. The urease activities were lower in the
cells grown on casamino acids throughout the life of the
culture. When urea was given in addition to casamino acids the
activities of the cells were similarly lower than the activities
of cells grown on urea alone. When cells were continuously
cultured on casamino acids the urease levels dropped quickly;
at the exponential phase of the 4th transfer period these
cells had a specific activity of 0.47. The casamino acids
present in the cell extracts did not inhibit the activity of

urease because dialysis of the extract from cells grown on

92

Figure 17. Increase in urease activity when cells
previously grown on ammonium succinate are

transferred back to urea medium.

Cells previously grown on ammonium succinate (from
experiment of Figure 16) were transferred to urea or
ammonium succinate medium and maintained on these media
for 5 transfer periods. At the exponential phase of
growth, the cultures were assayed for urease. Urease

:- formed/hr./gr.fr.wt.

for the cells grown on urea (o) and ammonium succinate (:3).

activity is expressed in umoles NH

93

 

 

 

 

 

:3 5.23 .E 3253 wzz 3.06.: 362:

2 4
Transfer

94

Figure 18- Decrease in urease activity in urea-conditioned

cells transferred to casamino acids medium.

Urea-conditioned cells in the stationary phase of growth
were transferred to urea or casamino acids medium. The cells
were assayed for urease at various times during the culture
+formed/

4
hr./gr.fr.wt. for the cells grown on urea (o) and casamino

period. Urease activity is expressed in umoles NH

acids (0).

95

 

 

 

 

l0

Culture Age (Days)

 

 

5. O 5.

A..3..:.._o\ of ooane ~12

3.25.: «moot:

96

casamino acids did not increase activity, and mixing of extracts
from cells grown on urea and cells grown on casamino acids did
notinhibit activity.

The above results indicate that the high levels of urease
fround in urea-conditioned cells are not stable and will
decrease to lower levels when the cells are cultured on a
different nitrogen source. The effect of nitrate on the
gradual decrease in urease activity was different than the other
nitrogen sources tested because in the presence of nitrate the
activity did not drop, even after 19 transfers, to the original

lower levels.

The Effects of Threonine on the Growth of Cells on Urea and

 

Other Nitrogen Sources

 

Threonine at a concentration of 100 uM inhibits growth
almost completely when XD cells are grown on nitrate as the
sole source of nitrogen (51). Heimer (80) showed that threonine
at the same concentration did not inhibit growth of urea-
adapted cells (previously grown on urea for at least one
transfer period) on urea. However, the work of Behrend and
Mateles (12) with cultured tobacco cells of the same XD
strain, but on a medium with a composition somewhat different
than M-lD, showed that 1 mM threonine inhibited growth on urea
(urea-adapted cells) and nitrate but not on ammonium succinate.
These authors suggested that the pathway for ammonium assimila-

tion was inhibited by threonine in cells grown on nitrate and

97

urea but not in cells grown on ammonium succinate. It was
suggested that the route for assimilation of endogenous ammonium
(cells grown on nitrate or urea) was different than for
exogenous ammonium (cells grown on ammonium succinate). However,
the 13N experiments reported in this thesis indicate that the
major pathway of ammonium assimilation is similar regardless of
the nitrogen source.

Because the effect of threonine on nitrate reductase and
nitrate uptake in the XD cells has been well documented (51,80),
an alternative explanation for the inhibitions of growth on urea
by the higher concentration of threonine reported by Behrend and
Mateles could be that threonine exerts some control over urea
utilization, but not necessarily through the same mechanism of
its effect on nitrate utilization. The experiments of Behrend
and Mateles differed from those of Heimer in the absolute
concentration of threonine used. In an attempt to resolve
whether the differences in the results obtained in these two
reports were due to the concentrations of threonine used, the
effects of different concentrations of threonine on growth on
different nitrogen sources was re-examined (Table V). The
results of Heimer were confirmed: 100 uM threonine did not
inhibit growth on urea. Higher concentrations of threonine
(300 uM and 1 mM) inhibited growth on urea, as Behrend and
Mateles reported. However, these higher concentrations of
threonine also inhibited growth on ammonium succinate. These

results suggest that the higher concentrations of threonine do

98

Table V. Effect of threonine on growth of cells maintained on

. . . a
nitrate, urea or ammonium succ1nate .

 

 

Medium grams fresh weight/l %
Nitrate 23.1 100
Nitrate + 100 uM threonine 0.6 2.6
Urea 19.4 100
Urea + 100 uM threonine 16.8 86.4
Urea + 300 uM threonine 1.3 6.6
Urea + 1 mM threonine 0.0 0.0
Ammonium succinate 18.6 100
Am. suc. + 100 uM threonine 14.4 77.4
Am. suc. + 300 uM threonine 0.0 0.0
Am. suc. + 1 mM threonine 0.0 0.0

 

aStationary phase cells maintained on nitrate, urea, or ammonium
succinate were subcultured into the same media :_threonine at
different concentrations. After 12 days the cells were
harvested and the fresh weight determined.

99

not uniquely effect growth on urea. The mechanism of
threonine inhibition at higher concentrations on all the
nitrogen sources tested could be due to its inhibitory effect
on the synthesis of the aspartate family amino acids (28).
Threonine does weakly inhibit the activity of urease in vitro
(Table VI), but only when the concentrations of threonine in
the reaction mixture are 5 and 10 times higher than that of
urea. Arginine, lysine and casamino acids at similar

concentrations did not inhibit urease activity in vitro.

Effect of Threonine on Growth of Cells which are Newly

Transferred from Nitrate to Urea or Ammonium Succinate Medium

 

Heimer (80) reported that growth of cells adapted to urea
was not inhibited by 100 uM threonine. However, cells which
were previously grown on nitrate and transferred to urea +
100 uM threonine did not grow. Although Heimer stated that
the cells needed to be cultured on urea for only one transfer
period before their sensitivity to threonine was totally lost,
cells may remain sensitive for a longer time. Table VII shows
the results of a growth experiment in which cells in the
stationary phase which were previously grown on nitrate were
transferred to nitrate, urea or ammonium succinate medium :
100 uM threonine. The cells were continuously grown on the
new nitrogen source, i_threonine, for 4 subsequent transfers
(3 for the ammonium succinate cells). Threonine inhibited

growth on all three nitrogen sources for the first transfer

100

Table VI. Effect of threonine on urease activity in vitroq

 

 

Threonine (mM) Ureaseb %
0 1.17 100

0.1 1.23 105.1

0.5 1.21 103.4

1.0 1.19 101.7

5.0 0.85 72.6

10.0 0.76 64.9

 

aCells grown on urea for 7 days were extracted and the extract
was dialyzed for 3 hours. The activity of urease was
determined in the presence of 1 mM urea and various concentra-
tions of threonine.

+

bumoles NH4

formed/hr./gr.fr.wt.

101

 

 

Table VII. Effect of threonine on growth of cells transferred
from nitrate medium to nitrate, urea or ammonium
. . a
succ1nate medium.
Transfer:
1 2
Medium __77_’—-—-
wt % wt % wt % wt %
Nitrate 28.4 100 26.3 100 24.5 100 25.0 100
Nit. + thr 0.1 0.3 19.6 74.5 11.8 48.2 6.1 24.4
Urea 11.6 40.8 13.5 51.3 13.6 55.5 14.2 56.8
Urea + thr 0.1 0.3 7.7 29.2 4.7 19.2 12.0 48.0
Am. suc. 18.7 65.8 18.3 69.6 17.3 70.6 -—- -—-
AS + thr 0.3 1.1 16.3 62.0 15.3 62.4 ——- '——

 

“Stationary phase cells maintained on nitrate were cultured into
medium containing nitrate, urea or ammonium succinate : 100 uM
After this first transfer period the cells growing
on each nitrogen source were subsequently cultured into medium
containing the same nitrogen source :_100 uM threonine.
13-14 days, at the end of each transfer period, the cultures

threonine.

(grown in triplicate) were harvested and the fresh weight
determined.

bgrams fresh weight/l.

After

102

period. Growth of cells on ammonium succinate was only slightly
inhibited by threonine for the 2nd and 3rd transfers. Growth on
urea was inhibited appreciably until the 4th transfer, i.e., the
cells retained their sensitivity to threonine during the course
of three transfers on urea. Cells grown on nitrate showed an
unexpectedly large variation in sensitivity to threonine during
these transfers, but the cells did remain sensitive in

agreement with many years of previous experience with these
cells.

Because nitrate cells retain their sensitivity to threonine
when transferred to urea medium, it was of interest to see how
long urea-conditioned cells would retain their resistance to
threonine inhibition when they were transferred to nitrate or
ammonium succinate medium. Table VIII shows the effects of
threonine (100 uM) on growth of cells transferred from urea to
medium containing one of the three nitrogen sources. Threonine
had very little effect on growth regardless of the nitrogen
source for the first transfer period. However, when the cells
which grew on nitrate + threonine medium were transferred to
the same medium, growth was dramatically inhibited. The cells
might have retained their resistance to threonine longer if the
cells grown on nitrate had been tested for threonine resistance
on subsequent transfers.

Cells grown on nitrate are physiologically different from
urea conditioned cells with respect to their sensitivity to

threonine. The loss of sensitivity to threonine when cells are

103

Table VIII. Effect of threonine on growth of cells transferred
from urea medium to urea, nitrate or ammonium

O O a
suCCinate medium .

 

 

 

Transfer:

Medium b 1' 2

wt % wt %
Urea 21.6 100 26.0 100
Urea + thr 22.3 103.2 22.9 88.1
Nitrate 19.8 91.6 18.6 71.5
Nit. + thr 20.1 93.1 0.1 0.4
Am. suc. 19.8 91.7 21.8 83.8
AS + thr 17.7 81.9 13.3 51.1

 

aStationary phase cells maintained on urea were cultured into
urea, ammonium succinate or nitrate medium :_100 uM threonine.
After 12 days the cells from each treatment were cultured into
the same medium and the remaining cells were harvested and the
fresh weight determined. All entries are averages of triplicate

samples.

bgrams fresh weight/l.

104

transferred from nitrate to urea occurs far more rapidly than
the gradual rise in urease activity observed in these cells,

suggesting that they are different phenomena. However, this

does not rule out the possibility that these two unusual

adaptations may have some common elements.

Effect of Tungstate on Growth of Cells

 

Sodium tungstate (100 uM) inhibits the formation of active
nitrate reductase in XD cells (80,83) by competing with
molybdenum. If nitrate reductase is the only enzyme in these
cells which requires molybdenum, then growth of cells on urea
or ammonium succinate should not be inhibited by tungstate.
When stationary phase cells maintained on nitrate, urea or
ammonium succinate were subcultured into the same media 1
100 uM sodium tungstate, growth was inhibited on all three
nitrogen sources (Table IX). The inhibition of growth does
not appear to be due to the competitive inhibition of ATP
sulfurylase by tungstate (145) because when 100 uM cysteine is
given as a supplemental sulfur source growth on urea is still
inhibited (Table IX). These results suggest that there may be
other enzymes which require molybdenum that are critically

important for growth of these cultured tobacco cells.

Search for the Presence of Urea Amidolyase in the Cultured

Tobacco Cells

 

Although there is no evidence for the presence of the

urea carboxylase/allophanate hydrolase system for urea

105

Table Ix. Effect of tungstate on growth of cells maintained

0 O I a
continuously on nitrate, urea or ammonium succ1nate.

 

 

Medium grams fresh weight/1 %
Nitrate 21.0 100
Nitrate + tungstate 0.0 0
Urea 19.0 100
Urea + tungstate 0.2 1.1
Urea + tungstate + cysteine 0.2 1.1
Urea + cysteine 19.1 100.5
Ammonium succinate 24.6 100
Ammonium succinate + tungstate 1.6 6.5

 

aStationary phase cells maintained on nitrate, urea or ammonium
succinate were subcultured into the same media :_100 uM sodium
tungstate. After 11 days of growth the cells were harvested
and the fresh weight determined. The effect of 100 uM cysteine

on growth on urea was also tested.r

106

hyrolysis in higher plants (148,177), an attempt was made to
determine if the cofactors required for this system would
stimulate urea hydrolysis in extracts of urea-conditioned cells.
The formation of ammonium was measured in dialyzed extracts
with or without the following additions: ATP, bicarbonate,
magnesium ions and potassium ions (Table X). A stimulation of
urea hydrolysis was detected with the addition of the above
cofactors, however avidin did not inhibit this stimulation.
Table X indicates that enhancement of activity was not
attributable to the presence of ATP but was due to the combined
presence of Kt. Mg++ and bicarbonate. When extracts of urea
conditioned cells are dialyzed, a decrease in urease activity
occurs. It was found that these compounds would stimulate the
activity of dialyzed extracts up to but not beyond the activity
present in the undialyzed extract. When the cations and
bicarbonate were added to undialyzed extracts no stimulation of
urea hydrolysis was observed. The stimulation of urease
activity by these compounds is probably due to a nonspecific
effect of replacing an ionic environment which was lost as a
result of the dialysis procedure. The above results suggest
that there is no urea amidolyase activity in extracts of

tobacco cells grown on urea.

Effect of Nickel and Potassium Citrate on Growth of Cells
Polacco (138) has reported that 10 mM potassium citrate

inhibits growth of soybean cells cultured on urea, but that

107

Table X. Test for stimulation of urea hydrolysis in presence of

 

 

ATPa.

Additions Ureaseb %
Urea only 0.73 100
Reaction mixturec 1.06 145
Reaction mix. + 200 ug avidin 1.11 152
Reaction mix. + 100 ug biotin 1.15 158
Reactsgnugizio:iioo pg aVidin 1.12 153
Mg“+ alone 0.65 89
143* + 15' 0.97 133
MS‘ + bicarbonate 1.03 141
K+ alone 0.93 127
K+ + bicarbonate 1.20 164
bicarbonate alone 0.97 133
Mg++ + x“ + bicarbonate 1.31 179

 

aCells grown on urea for 10 days were extracted for urease. The
extract was dialyzed for 2 hours.

4.
4

cThe reaction mixture contained 50 mM urea, 10 mM ATP, 10 mM
MgS04, 10 mM sodium bicarbonate and 40 mM KCl.

bumoles NH formed/hr./gr.fr.wt.

108

growth on other nitrogen sources is not inhibited by this
compound, and that the inhibition of growth on urea is alleviated

by the addition of 10 UM NiSO The effects of potassium citrate

4.
and NiSO4 on growth of tobacco cells on urea were examined

(Table XI). The addition of 10 uM NiSO4 to the medium did not
stimulate growth on urea. Growth on urea was inhibited by 3 and
10 mM potassium citrate, but 10 uM NiSO4 did not alleviate these
inhibitions. Cells grown on ammonium succinate were also
inhibited by 3 and 10 mM potassium citrate, but growth on
nitrate was only inhibited at the higher potassium citrate
concentration. The results indicate that nickel does not
stimulate growth on urea in the prsence or absence of potassium

citrate, and that potassium citrate does not inhibit growth

specifically on urea.

109

Table XI. Effects of potassium citrate and NiSO on growth of

4
. . . a
cells on urea, nitrate or ammonium succ1nate.

 

 

Medium grams fresh weight/1 %
Urea 30.2 100
Urea + 10 UM NiSO4 28.4 94.0
Urea + Ni + 1 mM K citrate 24.0 79.5
Urea + Ni + 3 mM K citrate 0.1 0.3
Urea + Ni + 10 mM K citrate 0.0 0.0
Urea + 1 mM K citrate 22.2 73.5
Urea + 3 mM K citrate 0.2 0.7
Urea + 10 mM K citrate 0.0 0.0
Nitrate 32.8 100
Nitrate + 3 mM K citrate 26.4 80.5
Nitrate + 10 mM K citrate 0.0 0.0
Ammonium succinate 24.1 100
Am. suc. + 3 mM K citrate 0.2 0.8
Am. suc. + 10 mM K citrate 0.0 0.0

 

“Stationary phase cells maintained on urea were cultured into
urea medium with various additions of NiSO and potassium
citrate. Stationary phase cells maintaine on nitrate or
ammonium succinate were transferred to the same medium.:_
various additions of potassium citrate. After 12 days the
cells were harvested and the fresh weight determined. A11
fresh weight entries are the aVerages of duplicate samples.

DISCUSSION

Urea is an effective source of nitrogen for plant growth.
In this regard, the X0 line of tobacco cells is no exception.
These cells grow on urea as a sole source of nitrogen, although
growth on nitrate is slightly better (Figure 1). The cells
contain urease (Figure 2), but do not contain detectable
amounts of the urea carboxylase/allophanate hydrolase enzymes
(Table X). When grown on urea, the highest levels of urea
accumulate in cells with the lowest urease activity (Figure 8),
and when the urease levels eventually increase in these cells
their urea concentrations drop. Cells with higher levels of
urease grow slightly faster on urea than cells exhibiting
lower levels of urease (Figure 12). These observations suggest
that urease is responsible for urea utilization in these cells.
The apparent induction of nitrate reductase (51) and of nitrite
reductase (203) by nitrate or nitrite, and the apparent de-
pression of acid phosphatase by the absence of phosphate (204)
and of ATP sulfurylase by the absence of sulfate (145) has
been documented in the XD cell line. The object of this
investigation was to study the regulation of urease in the

cultured tobacco cells. It was assumed that urea might

110

lll

similarly induce urease in these cells because of the previous
reports of its apparent induction in other higher plant tissues
and because it serves as an optional nitrogen source. Cells
grown continuously on urea for many generations have activities
of urease which are 4 to 5 times higher than those found in the
cells grown on nitrate (Figure 2). The results of kinetic
studies (Figures 4 and 5) suggest that the enzymes extracted
from cells grown on the two different nitrogen sources are
similar, in that they possess exactly the same Km for urea.

The urea-urease system differs from the classic nitrate-nitrate
reductase system in these cells. Cells which are newly
transferred to urea medium do not respond by making higher
levels of urease (Figure 7) as they respond by making nitrate
reductase when they are newly transferred to nitrate medium (51%
Cells which possess the higher levels of urease still retain
high levels at the stationary phase of growth, even if they are
transferred into freah medium containing nitrate and no urea
(Figure 13). Cells in the stationary phase of growth on
nitrate do not continue to produce nitrate reductase. Also,
cells in the exponential phase of growth on urea do not produce
nitrate reductase, while cells in the exponential phase of
growth on nitrate produce low levels of urease. The mechanism
determining the changes in the levels of urease in these cells
appears to be fundamentally different from that responsible for
the much faster changes associated with nitrate reductase and

other enzymes in these cells.

112

The discovery that there is no immediate response by the
tobacco cells to the presence of urea in the medium is
contradictory to what has been reported in rice plants (119),
potato plants (126), duckweed (20), and soybean tissue cultures
(137). Of these studies, the system for which the best
comparison can be made to that of the tobacco cell culture
system is the soybean cell culture system. When compared on a
per mg protein basis, the levels of urease found in soybean
cells (137) cultured on urea are very similar to those found in
tobacco cells which have been cultured on urea for many gener-
ations. The levels of urease found in the soybean cells grown
on a Murashige and Skoog (MS) medium were 1/2 to 1/4 of those
which are found in the tobacco cells grown on nitrate or
ammonium. The major difference between the ureases in these
two systems is that when the soybean cells are transferred from
MS medium to urea medium the urease activity rises to the
higher levels. This is not the case in the tobacco cells, in
which the urease levels rise only after numerous transfers on
urea (Figures 9 and 10). It is perplexing that in one system
(soybean) the enzyme is apparently inducible while in the
other system (tobacco) the enzyme appears to be constitutive
at two different levels.

It is unlikely that the gradual increase in urease activity
observed in cells grown on urea for many generations is due to
a selection of a spontaneous variant cell. The cells grow

when transferred from nitrate to urea, and it would take

113

considerably longer than the actual time observed for a
variant cell possessing a high activity of urease to take over
the cell population and exhibit a high urease activity in the
culture.

If the urease levels of the individual cells in the nitrate
cell population varied, e.g., if 1 in 10 cells had the high
urease, then the increase in urease could be attributed to a
gradual takeover of the cell population by these high-urease
cells. The cloning experiment confirms that this is not the
case (Figure 11). Although the urease levels in individual
clones from the population of cells grown on nitrate varied,
probability calculations based on the observed variation in
the population show that it is highly unlikely that any cells
in this population possess levels of urease around the specific
activity of 2.0. Similar calculations show that about 1 cell
in 33,000 in the population could have a urease activity of
1.0. It was calculated (using the growth rate constants from
Figure 12) that it would take 350 days for these cells to take
over the population and result in the expression of a specific
activity of 1.0. The actual observed times in which the
specific activity rose to 2.0 were 120 and 220 days after cells
grown on nitrate were transferred to and subsequently maintained
on urea. These results indicate that the increase in urease
activity cannot be due to a selection of a pre-existing

subpopulation of high-urease cells.

114

An alternate interpretation for this phenomenon is that all
or most of the cells slowly adapt to urea by making the higher
levels of urease. Evidence in favor of a slow adaptation was
provided when clones isolated from a population of cells on
nitrate and cultured on urea increased their urease levels after
10 transfers on urea (Table III).

When one considers mechanisms to explain the gradual
adaptation observed in the cultured tobacco cells, the classical
terms of induction or repression and derepression cannot be used
because of the differences in the time scales involved. The
apparent induction and repression processes described in higher
plants usually occur in hours or at most days. In the case of
urease in the tobacco pell system it takes months before any
change in enzyme levels are observed and weeks after that
before the higher levels are fully expressed. It is hard to
imagine an induction or derepression occurring here because it
would mean that the gene or genes for urease could gradually
produce different amounts of the enzyme, and that this variable
gene expression is continually transferred from generation to
generation. In other words, this is not an all or none
phenomenon like induction or derepression but instead is a
gradual process occurring over an extended period of time.

There are striking similarities between the increase in
dihydrofolate reductase levels observed in murine Sarcoma cells
(53,75,95,99,112,135) and the increase observed in urease levels

in the tobacco cell cultures. In both cases the changes involve

115

the level of a specific enzyme, the changes occur gradually,
and once the change has occurred the new levels remain stable
if the cells are kept under the same culture conditions. When
the culture conditions are changed (omission of methotrexate
from the Sarcoma cell culture medium or omission of urea from
the tobacco cell culture medium, Figure 16), the enzyme levels
are unstable and gradually decrease to the original lower
levels. In some cases the levels of dihydrofolate reductase
stabilize at an elevated level in the absence of methotrexate
(53,130). In the tobacco cells the urease levels are
stabilized at a point between the high and low levels when the
cells are transferred back to nitrate medium (Figure 14). It is
only fair to point out the differences in these two systems.
The rises in dihydrofolate reductase occur as a result of a
progressively more stringent stepwise selection procedure in
which the cells are exposed to increasingly higher concentrations
of methotrexate and most of the cells are killed at each
concentration step. The increase in urease does not involve a
drastic selection procedure. The increases observed in
dihydrofolate reductase activity can be ZOO-fold higher than
the normal levels, and dihydrofolate reductase can comprise as
much as 6% of the total soluble protein (3). The highest
increase observed in urease levels was 7-fold. It was
therefore quite feasible to isolate dihydrofolate reductase
mRNA (101), and synthesize complementary cDNA (2) from the

methotexate resistant murine Sarcoma cell lines. As a result,

116

it was proven quite recently that the increase in dihydrofolate
reductase levels is due to a multiplication of genes responsible
for the synthesis of this enzyme. Although the magnitude of the
changes with urease make this system a less favorable subject
for investigation, there are redeeming characteristics in this
system that should be pointed out. Since the changes occur in
the dihydrofolate reductase system as a result of treatment with
an analog of folic acid which is not found in nature, this is
not a process which one would expect to find normally in these
cells. The changes in urease levels occur in the tobacco cells
as a result of a change (exposure to urea) which could occur

to plants in nature. Even though the increases are typically
only 4 to 5 fold, they enable the cells to utilize urea more
effectively (Figures 8 and 12). A cautionary note must be

made to the effect that cells in culture are in an environment
which is rather different from the natural environment of a
plant cell. It is therefore uncertain if such a phenomenon
could occur in a whole plant as it grows. The work of Durant
(43), in which heritable changes were induced in flax plants
supplied with different fertilizers, and the work of Highkin
(85), in which heritable changes were induced in pea plants
subjected to continuous temperature conditions, both suggest
that phenomena of a similar type may occur in whole plants.
Although the observations seem Lamarckian in nature, they may
reflect the operation of mechanisms based upon a multiplication

of specific pre-existing genes.

117

A gradual duplication of genes responsible for urease
synthesis may be the mechanism responsible for the gradual
increase in the activity of urease. This hypothesis was
elegantly proven for the dihydrofolate reductase system.
However, for the reasons described above it would be more
difficult to test for the urease system. Nevertheless, one can
speculate on how such a process could occur in these cells.

In bacteria, tandem duplication of specific genes is
quite common (4,63,76,87) but the additional genes do not
remain as a permanent part of the genome. Gene duplication
also occurs in phages (47), Gene duplications can occur in
Drosophila by two mechanisms: by unequal interchromosomal
crossing over, as occurs at the bar locus (26,162); or by
unequal exchanges between sister chromatids, i.e., intra-
chromosomal exchange (136). Random unequal crossing over and
intrachromosomal exchange are possible mechanisms by which
repeated DNA sequences evolve (158). It was suggested (2) that
a tandem duplication of the dihydrofolate reductase gene might
occur by either of these mechanisms. The sister chromatid
exchange mechanism was favored, however, because it has been
demonstrated to occur in a variety of organisms, including
plants (116,122,173). Either of these mechanisms could be
occurring in the cultured tobacco cells. At each cell division,
one of the progeny could receive slightly more genes for urease
while the other receives slightly less. If such a process is

randomly happening to a large number of the cells in the

118

population, over an extended period of time under conditions
where urea is present in the medium, the cells with more urease
genes would be slightly favored and higher levels of urease
would eventually be expressed by the culture. Cell populations
grown on nitrate have various levels of urease, but no high
levels (Figure 11). Since they are always grown on nitrate, a
cell which makes slightly higher levels has no advantage over the
other cells. Although the urease levels are constant for these
cultures, on a cell to cell basis the levels may always be
changing. The urease levels in clones of urea conditioned cells
also vary. Some clones have levels 1 1/2 to 2 times higher

than what is found in the normal cell population. With the
above suggested mechanisms the cultures can make even higher
levels of urease. The estimatiOns of how long it would take
cells with a certain level of urease to take over a population
were calculated assuming that the urease level in each cell

is fixed. This may be greatly at variance with the real
situation. Perhaps one should envision the urease levels of

the cells in a dynamic state which always changes slightly

upon cell division.

If the increase in urease is due to the above mechanisms,
then it may be possible to employ a selection procedure which
would result in the selection of cells which produce even
higher levels of urease. This could be done by exposing the
cells to inhibitors of urease, such as hydroxyurea (16) or

derivatives of hydroxamic acids (106). The selection should be

119

performed by gradually increasing the concentration of the
inhibitor, exactly the way the cells resistant to methotrexate
were obtained. Another possible method by which higher levels
of urease might be obtained, would be to isolate, grow, and
reisolate clones from cloned populations which already have a
high level of urease. The clone with the highest urease
activity would be recloned, and so forth, so that the cells
which have the highest tendency to increase in urease would be
chosen. Since the jack bean seed possesses large amounts of
urease, the molecular basis for the synthesis of urease in this
plant tissue should be determined. If urease synthesis in the
jack bean seed is accompanied by the action of a large number
of genes for urease, it would be interesting to speculate on
what kind of natural selection process was responsible for

the multiplication of such genes.

Another mechanism for selective multiplication of genes
is the generation of extrachromosomal copies. Examples of such
a process are: the amplification of ribosomal genes in
amphibian oocytes (27), and reverse transcription of specific
viral mRNA (8,174). With these mechanisms a large number of
specific genes are produced in a single step. This is
probably not the mechanism responsible for the increase in
urease because here the increase is gradual. Alternatively,
an increase in the number of urease genes might be achieved by

the retention of specific chromosomal fragments. Transferred

120

genetic elements are usually lost rapidly from host cells in
culture (35,121,159,197), but can be maintained indefinitely by
growth under appropriate selective conditions. The number of
genes for a particular enzyme can also be increased by the
production of aneuploid cell lines in culture in which an
additional chromosome, which contains that particular gene,
becomes a part of the genome (206). e.g. trisomics would have

1 1/2 times the activity found in the diploid cell.

The slow habituation of cultured tobacco cells to a
condition of requiring no exogenous growth factors (124) bears
some resemblance to the slow adaptation of urease described in
this thesis. Cloning experiments showed that habituation occurs
in all the cells in the culture and that on the average the
cell population gradually becomes more and more habituated
(124). Similarly, the cloning experiments reported in this
thesis suggest that each cell has the potential to adapt to
making the higher levels of urease. The adaptation described
here involves a specific enzyme whereas the habituation process
could conceivably involve the action of many enzymes. The
habituation process may, however, be a valuable model for cell
differentiation in culture.

This thesis has described a slow adaptation phenomenon
that occurs in the cells of a higher plant. The molecular
mechanism responsible for the gradual changes in urease is not

yet established, and certainly merits investigation.

BIBLIOGRAPHY

10.

11.

12.

13.

BIBLIOGRAPHY

Alexander, M., in Soil Nitrogen, (Bartholomew, W. V., Clark,
F. E., Eds.), PP. 307-343 (1965), Am. Soc. Agron., Inc.,
Madison, WI.

 

Alt, F. W., Kellems, R. E., Bertino, J. R., Schimke, R. T.,
J. Biol. Chem., 253, 1357-1370 (1978).

Alt, F. W., Kellems, R. E., Schimke, R. T., J. Biol. Chem.,
251, 3063-3074 (1976).

Anderson, R. P., Miller, C. G., Roth, J. R., J. Mol. Biol.
105, 201-218 (1976).

Arkin, H., Colton, R. R., Tables for Statisticians (1970),
Barnes & Noble, Inc., New York.

Arnon, D. 1., Soil Sci., 44, 91-121 (1937).
Bailey, C. J., Boulter, D., in Chemotaxonomy of the

Leguminosae, (Harborme, J. B., Boulter, D., Turner, B. L.,
Eds.). pp. 485-502 (1971), Academic Press, New York.

 

Baltimore, D., Nature, 226, 1209-1211 (1970).
Barnes, R. L., Nature, 184, 1944 (1959).

Bayley, J. M., King, J., Gamborg, 0. L., Planta,105, 15-24
(1972).

Beaumont, A. B., Larsinos, G. J., Piekenbrock, P., Nelson,
P. R., J. Agric. Res. 43, 559-567 (1931).

Behrend, J., Mateles, R. I., Plant Physiol., 56, 584-589
(1975).

Behrend, J., Mateles, R. I., Plant Physiol., 58, 510-512
(1976).

121

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

122

Biedler, J. L., Spangler, B. A., J. Cell Biol., 29, 117a
(1976).

Binns, A., Meins, F., Jr., Proc. Natl. Acad. Sci. USA, 18,
2660-2662 (1973).

Blakeley, R. L., Hinds, J. A., Kunge, H. E., Webb, E. C.,
Zerner, 8., Biochem. J., 8, 1991-2000 (1969).

Blakeley, R. L., Webb, E. C., Zerner, B., Biochem. J., 8,
1984-1990 (1969).

Blattler, D. P., Contaxis, C. C., Reithel, F. J., Nature,
216, 274-275 (1967).

Bollard, E. G., Symp. Soc. Exptl. Biol., 18, 304-329 (1959).

Bollard, E. G., Cook, A. R., Turner, N. A., Planta, 88, l-
12 (1968).

Boulter, D., Davis, 0. J., New Phytol., 82, 935-946 (1968).
Boynton,D., Ann. Rev. Plant Physiol., 8, 31-54 (1954).
Braun, A. C., Symp. Soc. Exp. Biol., ll, 132-142 (1957).

Braun, A. C., Proc. Natl. Acad. Sci. USA, 33, 344-349
(1958).

Braun, A. C., Proc. Natl. Acad. Sci. USA, 28, 932—938
(1959).

Bridges, c. 13., Sci. 8_3_, 210-211 (1936).

Brown, D. 0., Dawid, I. B., Sci. 160, 272-280 (1968).
Bryan, J. K., in Plant Biochemistry: (Bonner, J., Varner,
J. E., Eds.). pp- 525-560 (1976), Academic Press, New
York.

Castric, P. A., Levenberg, 8., Biochim. Biophys. Acta, 438,
574-583 (1976).

Chapman, H. D., Liebig, G. F., Soil Sci. Soc. Am. Proc.,
18, 276-282 (1952).

COOk’ A. R. ' Planta, _8_3-' 13-19 (1968) 0

Cook, A. R., Boulter, D., Phytochem., 8_313-3l9 (1964).

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.,

49.

SO.

51.

123

Court, M. N., Stephen, R. C., Waid, J., Nature, 194,
1263-1265 (1962).

Creeth, J. M., Nichol, L. W., Biochem. J., 22, 230-239
(1960).

Degnen, G. E., Miller, I. L., Eisenstadt, J. M., Adelberg,
E. A., Proc. Natl. Acad. Sci. USA, Z8, 2838-2842 (1976).

Delluva, A. M., Markley, K., Davies, R. E., Biochim.
Biophys. Acta, 151, 646-650 (1968).

Dilworth, M. F., Dure, L., III, Plant Physiol., 88, 698-702
(1978).

Dintzis, R. Z., Hastings, A. 8., Proc. Natl. Acad. Sci. USA,
88, 571-578 (1953).

Dixon, N. E., Gazzola, C., Blakeley, R. L., Zerner, B.,
J. Amer. Chem. Soc., 82, 4131-4133 (1975).

Dougall, D. K., Biochem. Biophys. Res. Commun., 88, 639-646
(1974).

Dougall, D. K., Verma, D. C., In Vitro, 88, 180-182 (1978).
Dounce, A. L., J. Biol. Chem., 888, 307-308 (1941).
Durrant, A., Heredity, 81, 27-61 (1962).

Durrant, A., Nature, 888, 1302-1304 (1962).

Dyson, W. H., Hall, R. H., Plant Physiol., 88, 616-621
(1972).

Einset, J. W., Skoog, F., Proc. Natl. Acad. Sci. USA,
28, 658-660 (1973).

Emmons, S. W., MacCosham, U., Baldwin, R. L., J. Mol. Biol.,
88, 133-146 (1975).

Fasman, G. D., Nieman, C., J. Amer. Chem. Soc., 18, 1646-
1650 (1951).

Filner, P., Ph.D. Thesis, California Institute of
Technology, Pasadena (1965).

Filner, P., Exp. Cell Res., 88, 33-39 (1965).

Filner, P., Biochim. Biophys. Acta, 118, 299-310 (1966).

124

52. Filner, P., Dev. Biol. Suppl., 8, 206-226 (1969).

53. Fischer, G. A., Biochem. Pharmacol., 1, 75-80 (1961).

54. Fishbein, W. N., Fifth International Symposium on
Chromatography and Electrophoresis (1969), Ann Arbor-
Humphrey Science Publishers, Inc., Ann Arbor, pp. 238-241.

55. Fishbein, W. N., J. Biol. Chem., 244, 1188-1193 (1969).

56. Fishbein, W. N., Engler, W. F., Griffin, J. L., Scurzi, W.,
Bahr, G. F., Eur, J. Biochem., 18, 185-190 (1977).

57. Fishbein, W. N., Nagarajan, K., Scurzi, W., Arch. Biochem.
Biophys., 172, 726-733 (1976).

58. Fishbein, W. N., Spears, C. L., Scurzi, W., Nature, 223,
191-193 (1969).

59. Fishbein, W. N., Winter, T. 8., Davidson, J. D., J. Biol.
Chem. 240, 2402-2406 (1965).

60. Fisher, E. G., Proc. Amer. Soc. Hort. Sci., 88, 91-98
(1952).

61. Fisher, E. G., Cook, J. E., Proc. Amer. Soc. Hort. Sci.
88, 35-40 (1950).

62. Fisher, W. 3., Parks, W. L., Proc. Soil Sci. Sci. Amer.
88, 247-248 (1958).

63. Folk, W. R., Berg, P., J. Mol. Biol., 88, 595-610 (1971).
64. Fox, J. E., Physiol. Plant., 88, 793-803 (1963).

65. Freiberg, S. R., Payne, P., Proc. Amer. Soc. Hort. Sci.,
88, 226-234 (1957).

66. Gamborg, 0. L., Plant Physiol., 88, 372-375 (1970).

67. Gamborg, 0. L., Miller, R. A., Ojima, K., Exp. Cell Res.,
88, 151-158 (1968).

68. Gamborg, 0. L., Shyluk, J. P., Plant Physiol., 88, 598-600
(1970).

69. Gorin, G., Biochim. Biophys. Acta, 88, 268-270 (1959).

70. Gorin, G., Chin, C.-C., wang, S. F., Experientia, 88, 685-
687 (1968).

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

88.

89.

125

Gorin, G., Fuchs, F., Butler, L. G., Chopra, 5. L., Hersh,
R. T., Biochem. 8, 911-916 (1962).

Granick, 5., Plant Physiol., 88, 471-486 (1937).
Granick, 5., Plant Physiol., 88, 601-623 (1937).
Granick, 5., Plant Physiol., 88, 29-54 (1937).

Hakala, M. T., Zakrazeuski, S. F., Nichol, C. A., J. Biol.
Chem., 236, 952-958 (1961).

Hariuchi, T., Hariuchi, S., Novick, A., Genetics, 88, 157-
169 (1963).

Hattori, A., J. Biochem., 88, 253-273 (1957).
Hattori, A., J. Biochem., 88, 57-64 (1958).
Hattori, A., Plant and Cell Physiol., 8, 107-115 (1960).

Heimer, Y. A., Ph.D. Thesis, Michigan State University,
East Lansing (1970).

Heimer, Y. M., Filner, P., Biochim. Biophys. Acta, 215,
152-165 (1970).

Heimer, Y. M., Filner, P., Biochim. Biophys. Acta, 230,
362-372 (1971).

Heimer, Y. M., Wray, J. L., Filner, P., Plant Physiol.,
82, 1197-1199 (1969).

Heywood, V. H., in Chemotaxonomy_of the Leguminosae,
(Harbone, J. B., Boulter, D., Turner, B. L., Eds.),
pp. 1-29 (1971), Academic Press, New York.

Highkin, H. R., Am.J. Bot., 52! 626-631 (1958).
Highkin, H. R., Nature, 182, 1460 (1958).

Hill, C. W., Combriato, G., Molec. Gen. Genet., 127, 197-
214 (1973).

Hinsvark, 0. N., Wittwer, S. H., Tukey, H. 8., Plant
Physiol., 88, 70-76 (1953).

Hirose, 5., Goto, Y., Soil Sci. and Plant Nutr., 7,
85 (1961).

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

106.

107.

126
Hoare, J. P., Laidler, K. J., J. Amer. Chem. Soc., 22,
2487-2489 (1950).

Hodson, R. C., Thompson, J. F., Plant Physiol., 88, 691-
696 (1969).

Hodson, R. C., Williams, S. K., Davidson, W. R., J.
Bacteriol., 121, 1022-1035 (1975).

Hughes, R. B., Katz, S. A., Stubbins, S. E., Enzymologia,
88, 332-334 (1969).

Jablonski, J. R., Skoog, F., Physiol. Plant., 2, 16-24
(1954).

Jackson, R. C., Huennekens, F. M., Arch. Biochem. BiOphys.,
154, 192-198 (1973).

Jones, R. W., Abbott, A. J., Hewitt, E. J., Jones, D. M.,
Best, G. R., Planta, 133, 27-34 (1976).

Jones, V. M., Boulter, D., New Phytol., 8] 925-934 (1968).
Kaplan, A., in Standard Methods of Clinical Chemistry

(Meites, 8., Ed.), Vol. 5, pp. 245-256 (1965), Academic
Press, London.

 

Kashet, E. R., Crawford, E. J., Friedkin, M., Humphreys,
S. R., Golding, A., Biochem. 8, 1928-1931 (1964).

Rating, H., Biochem. Z., 335, 366-381 (1962).

Kellems, R. E., Alt, F. W., Schimke, R. T., J. Biol. Chem.,
251, 6987-6993 (1976).

Kiesel, A., z. Physiol. Chem. 28, 169-196 (1911).

Kistiakowsky, G. B., Mangelsdorf, P. C., Rosenberg, A. J.,
Shaw, W. H. R., J. Amer. Chem. Soc., 28, 5015-5020 (1952).

Kistiakowsky, G. 8., Shaw, W. H. R., J. Amer. Chem. Soc.,
28, 866-871 (1953).

Kleczknowski, K., Cohen, P. P., Arch. Biochem. Biophys.,
107, 271-278 (1964).

Kobashi, K., Hase, J., Ushara, K., Biochim. Biophys. Acta,
88, 380-383 (1962).

Kollefel, C., Stroband, H. W. J., Phytochem., 88, 2635-
2638 (1973).

 

108.

109.

110.

111.

112.

113.

114.

115.

116.

117.

118.

119.

120.

121.

122.

123.

124.

127

Kollaffel, C., Van Dijke, H. D., Plant Physiol., 88, 507-
510 (1975).

Krebs, H. A., Henseleit, K. Z., Z. Physiol. Chem., 210,
33-66 (1932).

Kuykendall, J. R., Wallace, A., Proc. Am. Soc. Hort. Sci.,
88, 117-127 (1954).

Leftley, J. W., Syrett, P. J., J. Gen. Microbiol., 22,
109-115 (1973).

Littlefield, J. W., Proc. Natl. Acad. Sci. USA, 88, 88-95
(1969).

Lowan, A. N., Tables of Probability Functions, Vol. II
p. 340 (1942) Work projects administration for the city
of New York, sponsored by the National Bureau of
Standards.

 

Lowry, 0. H., Rosebrough, N. J., Farr, A. J., Randell,
R. J., J. Biol. Chem., 193, 265-275 (1951).

Maliga, P., Lazér, G., Svéb, Z., Nagy, F., Molec. Gen.
Genet., 149, 267-271 (1976).

Marin, G., Prescott, D. M., J. Cell Biol., 88, 159-167
(1964).

Mateer, J. G., Marshall, E. K., Jr., J. Biol. Chem., 88,
297-305 (1916).

Matsumoto, H., Hasegawa, Y., Kobayashi, M., Takahashi, E.,
Phys. Plant., 88, 872-881 (1968).

Matsumoto, H., Yasuda, T., Kobayashi, M., Takahashi, E.,
Soil Sci. and Plant Nutr., 82, 33-38 (1966).

Matsumoto, T., Yatazawa, M., Yamamoto, Y., Plant and
Cell Physiol., 88, 459-462 (1977).

McBride, 0. W., Ozer, H. L., Proc. Natl. Acad. Sci. USA,
28, 1258-1262 (1973).

McClintock, B., Cold Spring Harbor Symp. Quart. Biol., 8,
72-80 (1941).

Meins, F., Jr., Differentiation, 8, 21-25 (1973).

Meins, F., Jr., Binns, A., Proc. Natl. Acad. Sci. USA,
28, 2928-2932 (1977).

125.

126.

127.

128.

129.

130.

131.

132.

133.

134.

135.

136.

137.
138.
139.
140.

141.

142.

143.

128

Milton, J. M., Taylor, I. E. P., Biochem. J., 113, 678-
680 (1969).

Mokronosov, A. T., Ilinykh, z. G., Shukoloyukova, N. I.,
Fiziol. Rastenii, 88, 707-713 (1966).

Murashige, T., Skoog, F., Physiol. Plant., 88, 473-497
(1962).

Musculus, M., Compt. Rend. Acad. Sci., 28, 132-134 (1876).

Nabors, M. W., Daniels, A., Nadolny, L., Brown, C., Plant
Sci. Lett., 8, 155-159 (1975).

Nakamura, H., Littlefield, J. W., J. Biol. Chem., 247,
179—187 (1972).

Nakatsu, S., Haratake, S., Sakurai, Z., Zyo, N., Nishihara,
Z., Hayashida, M., J. Biochem., 8, 35-39 (1964).

Nanney, D. L., Proc. Natl. Acad. Sci. USA, 88, 712-717
(1958).

Oaks, A., Biochim. Biophys. Acta, 372, 122-126 (1974).

Pandita, M., Durzan, D. J., Proc. Can. Soc. Plant
Physiol., 88, 62 (1970).

Perkins, J. P., Hillcoat, B. L., Bertins, J. R., J. Biol.
Chem., 242, 4771-4776 (1967).

Peterson, H. M., Laughuan, J. R., Proc. Natl. Acad. Sci.
USA, 88, 126-133 (1963).

Polacco, J. C., Plant Physiol., 88, 350-357 (1976).

Polacco, J. C., Plant Physiol., 88, 827-830 (1977).

Polacco, J. C., Plant Sci. Lett. 88, 249-255 (1977).
Reinbothe, H., Flora, 888, 315-328 (1961).

Reinbothe, H., Mothes, K., Ann. Rev. Plant Physiol., 88,
129-150 (1962).

Reithel, F. J., in The Enzymes, (Boyer, P. D., Ed.), Vol.

 

Reithel, F. J., Robbins, J. E., Arch. Biochem. Biophys.
120, 158-164 (1967).

144.

145.

146.

147.

148.

149.
150.

151.

152.

153.

154.

155.

156.

157.

158.

159.

160.
161.
162.

163.

129

Reithel, F. J., Robbins, J. E., Gorin, G., Arch. Biochem.
Biophys., 108, 409-413 (1964).

Reuveny, Z., Ph.D. Thesis, Michigan State University,
East Lansing (1975).

Roon, R. J., Levenberg, B., J. Biol. Chem., 243, 5213-5215
(1968).

Roon, R. J., Levenberg, B., J. Biol. Chem., 245, 4593-4595
(1970).

Roon, R. J., Levenberg, B., J. Biol. Chem., 247, 4107-4113
(1972).

Rosenthal, G. A., Plant Physiol., 88, 273-276 (1970).
Rosenthal, G. A., J. Exp. Bot., 88, 609-613 (1974).

Sehgal, P. P., Naylor, A. W., Plant Physiol., 88, 567-572
(1966).

Shadaksharaswamy, M., Hill, R. M., Arch. Biochem. Biophys.
82, 607-609 (1962).

Shargool, P. D., FEBS Lett., 88, 348-350 (1973).

Shargool, P. D., Cossins, A., Can. J. Biochem., 82, 467-
475 (1969).

Shaw, W. H. R., J. Amer. Chem. Soc., 28, 2160-2163 (1954).

Shaw, W. H. R., Raval, D. N., J. Amer. Chem. Soc., 88,
3184-3187 (1961).

Siegel, L. M., Monty, K. J., Biochem. Biophys. Res.

Smith, G. P., Science 191, 528-535 (1976).

Spandidos, D. A., Siminovitch, L., Proc. Natl. Acad. Sci.
USA, 28, 3480-3484 (1977).

Splittstoesser, W. E., Phytochemistry, 8, 753-758 (1969).
Stern, C., Genetics, 88, 625-730 (1936).
Sturtevant, A. H., Genetics, 88, 117-147 (1925).

Sumner, J. B., J. Biol. Chem., 88, 435-441 (1926).

 

164.

165.

166.

167.

168.

169.

170.

171.

172.

173.

174.

175.

176.

177.

178.

179.

180.

181.

130

Sumner, J. B., J. Biol. Chem., 28, 97-98 (1926).
Sumner, J. B., Ergeb. Enzymforsch. 8, 295-301 (1932).
Sumner, J. B., in The Enzymes, (Sumner, J. B., Myrbach, K.,

Eds.), Vol. I, Part 2, pp. 873-892 (1951), Academic Press,
New York.

 

Sumner, J. B., Gralén, N., Quensel, I. B. E., J. Biol.Chem.,
125, 37-44 (1938).

Sumner, J. B., Hand, D. B., J. Biol. Chem., 28, 149-162
(1928).

Sumner, J. B., Hand, D. B., Holloway, R. G., J. Biol. Chem»
88,333-341 (1931).

Takeuchi, T., Coll. Agr. Tokyo Imp. Univ., 8, 1-14 (1909).

 

Tanisr R- J-: Naylor, A. W., Biochem. J., 108, 771-777
(1968).

Tartof, K. D., Ann. Rev. Genet., 8, 355-385 (1975).

Taylor, J. H., Woods, P. S., Hughs, W. L., Proc. Natl.
Acad. Sci. USA, 88, 122-128 (1957).

Temin, H., Mizutani, S. M., Nature, 226, 1211-1213 (1970).

Thomas, W. H., Krauss, R. W., Plant Physiol., 88, 113-122
(1955).

Thompson, J. F., Muenster, A. E., Biochem. Biophys. Res.
Commun., 88, 1049-1055 (1971).

Thompson, J. F., Muenster, A. E., in Mechanisms of
Regulation of Plant Growth, (Bieleski, R. L., Ferguson,
A. R., Cresswell, M. M., Eds.), Bulletin 12 (1974), The
Royal Society of New Zealand, Wellington.

 

 

Topfer, V. R., Mierach, J., Reinbothe, H., Biochem.
Physiol. Pflanzen., 161, 231-242 (1970).

Tschiersch, B., Flora, 147, 405-416 (1959).

Valmikinathan, K., Rao, V. N. V., Verghese, N.,
Enzymologia, 88, 257-268 (1968).

Van Slyke, D. D., Archibald, M., J. Biol. Chem., 154,
623-656 (1944).

182.

183.

184.

185.

186.

187.

188.

189.

190.

191.

192.

193.

194.

195.

196.

197.

198.

131

Van Slyke, D. D., Cullen, G. E., J. Biol. Chem., 88, 211-
228 (1914).

Varner, J. E., in The Enzymes, (Boyer, P. D., Lardy, H.,
Myrback, K., Eds.), Vol. 4, 2nd Edition, pp. 247-251
(1960), Academic Press, New York.

 

Viets, F. G., in Soil Nitrogen, (Bartholomew, W. V., Clark,
F. E., Eds.), pp. 503-549 (1965), Am. Soc. of Agronomy,
Inc., Madison, WI.

 

Waheed, A., Castui, P. C., J. Biol. Chem., 252, 1628-
1632 (1977).

Walker, J. B., Proc. Natl. Acad. Sci. USA, 88, 561-566
(1952).

Wall, M. C., Laidler, K. J., Arch. Biochem. Biophys. 88,
299-318 (1953).

Webster, G. C., Varner, J. E., Gansa, A. N., Plant

Weil, L., J. Franklin Inst., 238, 145 (1944).

Whiteside, J. A., Thurman, D. A., Planta, 88, 279-284
(1971).

Whitney, P. A., Cooper, T. G., Biochem. Biophys. Res.
Commun., 88, 814-819 (1970).

Whitney, P. A., Cooper, T. G., Biochem. Biophys. Res.
Commun., 88, 45-51 (1972).

Whitney, P. A., Cooper, T. G., J. Biol. Chem., 247,
1349-1353 (1972).

Whitney, P. A., Cooper, T. G., J. Biol. Chem., 248,
325-330 (1973).

Widholm, J. M., in Tissue Culture and Plant Science,
(Street, H. E., Ed.). pp. 287-289 (1974), Oxford.

 

Widholm, J. M., Can. J. Bot., 31, 1523-1529 (1976).

Willecke, K., Ruddle, F. H., Proc. Natl. Acad. Sci. USA,
28, 1792-1796 (1975).

Williams, W. T., Nature, 165, 79 (1950).

199.

200.

201.

202.

203.

204.

205.

206.

132

Williams, W. T., Sharma, P. Ch., J. Exp. Bot., 8, 136-
157 (1954).

Wood, H. N., Braun, A. C., Brandes, H., Kende, H., Proc.

Young, M., J. Exp. Bot., 88, 1172-1185 (1973).
Guitton, Y., Physiol. Veg., 8, 95-156 (1964).

Kelker, H. C., Filner, P., Biochem. Biophys. Acta, 252,
69-82 (1971).

Ueki, K., Sato, S., Physiol. Plant., 88, 506-511 (1971).

Drummond, M. H., Gordon, M. P., Nester, E. W., Chilton,
M. D., Nature, 269, 535-536 (1977).

Carlson, P. 5., Mol. Gen. Genet., 114, 273-280 (1972).

 

II. THE INITIAL ORGANIC PRODUCTS OF ASSIMILATION OF [13N]AMMONIUM

1
AND [ 3N]NITRATE BY TOBACCO CELLS IN CULTURE

INTRODUCTION

Ammonium assimilation in higher plants was long thought to
begin with the synthesis of glutamate by glutamate dehydrogenase.
However there is now reason to believe that the major route for
assimilation of ammonium may be the glutamine synthetase-
glutamate synthase pathway (21). Evidence for the presence of
this pathway in higher plants is largely based on enzymological
studies and on studies performed with 15N-labeled nitrogen.
Glutamine synthetase has been found in several plant tissues
(7,23,29) and was observed to be localized in the chloroplasts
(12,13,22). A pyridine nucleotide-dependent glutamate synthase
has been found in extracts of tissue cultures from six
different species (5,6) and a ferredoxin-dependent glutamate
synthase has been described from leaf tissue (15). However,
the detection of these activities in plant tissues does not
prove that they have a substantial role in the assimilation of
ammonium.

l

15 [ 5Nlamide glutamine and [1%flamino glutamate

When N05,
15 .
were fed separately to pea leaves, N appeared in free and

protein-bound amino acids in similar proportions regardless of

134

135

the labeled substrate applied (18). This observation was
consistent with assimilation via the glutamine synthetase-
glutamate synthase pathway because the 15N would presumably have
had to have passed sequentially through these compounds in

order for the amino acids to become labeled in similar propor-
tions. Studies done with a lO-hr feeding of 15Nogvia the F

transpiration stream of cut shoots of Dhtura demonstrated that

glutamine was the predominant recipient of photosynthetically

 

reduced nitrogen in the leaf (16,17). A kinetic study of the
5 .
appearance of 1 N in soluble nitrogenous compounds in roots of l

rice seedlings supplied with ISNHJ' or 15N05- showed that
glutamine was the most rapidly labeled compound, with con-
current but slower labeling of glutamic acid, aspartic acid and
alanine, in that order (32,33). The shortest times for which

the incorporation of label was tested was 5 min in the 15N0;'

studies and 15 min in the lsNHZ' studies. In a recent
. . . . . 5 - .
investigation, young pea leaves were supplied with 1 N0 in

3

the light and dark, and the incorporation of 15N into various
components of the leaves was followed (3). A large portion of
the 15N appeared in the amide group of glutamine and turned
over rapidly. Glutamic acid and alanine also became labeled,
although less than glutamine. Incorporation into aspartic
acid,ts-aminobutyric acid and homoserine was slower. The
shortest period of labeling in these studies was 1 hour.

The demonstration of the presence of both glutamine

synthetase and glutamate synthase in plants and the studies of

136

the kinetics of labeling are consistent with an involvement of
the glutamine synthetase-glutamate synthase pathway in the
assimilation of ammonium by higher plants. However, a precursor-
product relationship between the metabolites involved has not
been established; it has not been tested whether the assimilatory
pathway is dependent upon the nitrogen source used for growth;
and there has been only one instance (1) where use has been made
of inhibitors of the enzymes involved in the pathway, to test
whether the labeling observed is dependent on those enzymes.

The use of 13N in studies of nitrogen metabolism has been
limited because of the short half-life (10 min), the requirement
of a cyclotron and the lack of adequate procedures for prepar-
ation of the desired labeled compound. Ruben et al. briefly
tried utilizing 13N-labeled nitrogen gas to study N2 fixation
in non-leguminous plants (37). 099 used 13N to study the
mechanism of decomposition of nitrogen pentoxide (36).

Carangal and Varner bombarded thin graphite targets with protons
or deuterons in a cyclotron, digested them in acid mixtures and
prepared labeled nitrate, nitrite and ammonium compounds which
were then supplied to excised plant tissues (4). Nicholas

et al. (35) and Campbell et al. (34) reported preliminary
results using 13N-labeled nitrogen gas to study nitrogen
fixation in bacteria. In both studies the radioisotope was
taken up and incorporated into the bacterial cells. In all of
the above studies of nitrogen fixation the chemical problems

involved with the production of 13N-labeled nitrogen gas were

 

137

not solved. It was not possible to make [13N]N2 (only one atom

of the molecule is radioactive), therefore the counts may have

been incorporated into the organisms by reactions with 13N-

labeled free radicals which are produced when one of the

13N_13

radioactive atoms of a' N molecule decays. More recently

Wolk et al. reported the production of [13N1N2 by proton
bombardment of 13C and Dumas combustion of the target (30).
This method resulted in higher yields of 13N making studies of
nitrogen metabolism more feasible. 13N fixed by algal
filaments was localized by an autoradiographic technique.
Methods were later developed to elucidate the pathway of

assimilation of NZ-derived nitrogen using [13N]N2 (25,31). It

was demonstrated that Nz-derived nitrogen fixed by Anabaena

cylindrica is metabolized initiallylfirthe glutamine synthetase-

glutamate synthase pathway. Additional methods were developed

to produce 13NHIand it was shown that A. cylindrica

assimilates ammonium mainly via the same pathway (19,26).

13N05 has been used to determine rates of denitrification in

soils (11,27).

In this paper, we report on the initial products of

assimilation of 13NH2' by tobacco cells cultured with nitrate,

urea, or ammonium succinate as the sole nitrogen source, and

of assimilation of 13N03 by tobacco cells grown with nitrate

3

by proton bombardment of water differed from earlier methods

. 13
as the sole nitrogen source. The method of produCing N0

 

 

138

(11.27.28) in that 18O-depleted water was used as the target.
The production of 18F (t15 = 109.7 min) was thereby reduced by
approximately 90%. By studying assimilation for shorter times
than has heretofore been practicable, and by combining studies
of isotopic labeling with studies of the effects of inhibitors,
we have produced clear evidence that the glutamine synthetase-
glutamate synthase pathway is the most important, but, on the
other hand, not the sole, pathway for assimilation of inorganic

nitrogen by cultured cells of a higher plant.

 

 

MATERIALS AND METHODS

The Tobacco Cell Line and Growth Conditions
The tobacco cell line, growth media and growth conditions
were as described in the MATERIALS AND METHODS section of

Part I.

. + -
Generation oanH‘ and 13NO,
F1 .3

+
13N for production of 13NH was generated by irradiation of

4
18.6 mg of 13C (97 atom %, Monsanto Research Corp., Mound
Laboratory, Miamisburg, OH) with protons (30,2). The nuclear
reaction involved is 13C(p,n)13N. Beams of 12 MeV protons from
the Michigan State University cyclotron ranged from 0.7 to
4.0 uA, and bombardments lasted for 20 min. 13NHIwas produced
by Kjeldahl digestion by Thomas‘ modification (26) of the
method of Carangal and Varner (4). The target was transferred to
a 100-ml Kjeldahl flask which was preheated to 100° C on a
heating mantle. Potassium dichromate, 0.04 g, and 0.2 g of
potassium sulfate were added to the flask and mixed well with
the target, and 2 m1 of concentrated sulfuric acid (specific

gravity, 1.84) saturated with potassium dichromate was then

added. The temperature in the flask rose steadily reaching

139

 

 

140

nearly 250° C at the end of 10 min. The flask was then removed
from the heating mantle, and 5 m1 of saturated sodium borate
(pH 10), 8 ml of 40% w/v NaOH and 2 m1 of saturated silver
sulfate solution were added in rapid succession. lBNHJ. was
separated from the reaction mixture by distillation under
vacuum at 50° C. A side-arm test tube immersed in liquid r‘
nitrogen served as a receiving tube and radioactivity was

monitored with a model ABG-lOKG-SB ionization gauge (Jordon

Electronics, Division of Victoreen Instruments Co., Alhambra, CAL

 

Bumping and "boiling-over" of the reaction mixture was prevented b;

by permitting a feeble stream of air to enter the distillation

apparatus. The procedure for preparation of 13NH+. took

4
. . . +
approximately 15 min. The resultant frozen solution of 13NH4 ,

which contained an average of 6 mCi of 13NH;'

0.3 umol of NH+, was thawed and diluted to 1-2 ml. Samples of

and approximately

25 to 100 ul were transferred to a scintillation vial, 10 m1 of
"Cocktail D" (Beckman Instruments, Irvine, CA) was added, and
the radioactivity was measured with a Beckman CPM-100
scintillation counter. All measurements of radioactivity were

corrected for background and for time of decay.

13NOS' was generated by the 16O(P.a)13N reaction with

water (27). One ml of water, 99.98 atom.percent 160 containing
15 ppm 180 (Monsanto Research Corporation, Mound Laboratory)
was bombarded for 10 min with the same beam of protons described

above. The resultant radioactive solution, which contained

and 13NH4' in addition to 13N0"

4 3, was

varying amounts of 13N05'

 

141

subjected to the following purification procedure. The sample

was first oxidized using 1% H

13 -' 13 -
N02 to N03.

202, pH 2, for two min to convert
To remove l3NHJ, the pH was raised to 10 and
the solution evacuated to dryness. The residue was dissolved

in 0.5 m1 of H20 containing 10 mM 2(N-morpholino)ethane sulfonic
acid (MES) buffer (Sigma Chemical Co., St. Louis, MO), pH 6.0.
These purification procedures took approximately 10 min. The
resultant solution contained an average of 6 mCi of 13N0;:
Samples of 10 to 50 ul were taken for measurement of radioactivity
by liquid scintillation spectrometry. Purity was assessed in two
ways: a) Samples of 5 ul were taken before and after the
purification procedure and spotted on an area (2 x 10 mm) of a
glass plate (5 x 20 cm) coated with a 0.1 mm layer of cellulose
(E. Merck, Darmstadt, W. Germany). The plates were sprayed with
70 mM formic acid, pH 2.0, and subjected to electrophoresis at
3000 V for 2 min in a High Voltage Electrophoresis Apparatus,
model Qll-SAE-3203 (Shandon Scientific Co., London, England).
After electrophoresis, the plates were scanned for radioactivity
with a model 7201 Radiochromatogram Scanner (Packard Instrument
Co., Downers Grove, IL), using a 2.5 mm wide slit. Two peaks

of radioactivity were detected after electrophoresis and

scanning. One peak represented 13NO- and 13N052 while the

3
+ . . .
other peak corresponded to 13NH4. RadioactiVity was determined
13
by integration of the peak areas, and the percent NH4

determined after correction for the time interval between peaks.

b) Alternatively, samples of 10 ul were taken before and after

 

 

142

the purification procedure, and radioactive constituents were
separated by high performance liquid chromatography (HPLC)
using a Partisil 10 SAX (Whatman Inc., Clifton, NJ) anion
exchange column (28) equipped with an Instrument Mini-Pump

(Milton Roy Co., Riviera Beach, FL). The elution buffer was

50 mM phosphate, pH 3.0. Fractions corresponding to 13NH+

4 I
13 - 13 - . . . .
N02, and N03 were eluted separately, and their radioactiVity

measured with a sodium iodide coincidence counter. The
radioactivity in each peak was integrated, and the percent 13N
in each constituent determined. In one experiment, 13nog' was
purified preparatively by HPLC.

When expressed as a percent of total 13N, the unpurified

sample of 13:40; contained 2.0 i 1.4% 131411; and 6.1 i 3.6% 131102”.

After oxidation with hydrogen peroxide and evaporation to dryness

at alkaline pH these contaminations decreased to 0.1 :_0.1%

13NH;. and 0.8 :_0.5% 13NO£1 All 13N0;' experiments were
performed with purified samples. The ratio of radioactivity of
18

F to 13N in solution at the time of generation of 13N0;' was

approximately 3 x 10-5

Assimilation of 13NH:' and 13NO§8

 

In preparation for the assimilation of 13N, cells from
4-7 day old cultures in the exponential phase of growth were
harvested by vacuum filtration on Whatman No. 1 filter paper.
Twenty mg fresh weight of cells per sample were resuspended

in a small volume (0.2-0.4 ml, depending upon the experiment) of

 

143

0.1 mM sodium succinate, pH 6.2, in lZ-ml conical centrifuge

tubes. In some experiments, 1 mM L-methionine-DL-sulfoximine
(Sigma Chemical Co., St. Louis, MO) or O-diazoacetyl-L-serine
(azaserine: Calbiochem, La Jolla, CA) was added to the cell

suspension. The tubes containing suspensions of cells and

13

N-labeled substrate were swirled continuously or periodically

by means of a Super-Mixer (Arthur H. Thomas Co., Philadelphia,

PA). The cells were exposed to 13NH;' for periods of 10 s to
3 _
10 min and to 1 NO for periods of 30 s to 15 min. The

3

assimilation of 13N was stopped by mixing the cell suspensionwdth
four volumes of 100% methanol. The cells were removed by
centrifugation at 1000 x g for 2 min. The radioactive organic
products in the methanolic extracts were concentrated;

separated by high-voltage electrophoresis with 70 mM borate
buffer, pH 9.2, on plates coated with thin layers of cellulose
(0.1 mm); localized and quantified by radiochromatogram

scanning; and identified tentatively by coelectrophoresis with
stable amino acid standards at pH 9.2 (19,25,31). The elapsed
time from the end of bombardment to final counting ranged from

1 to 2.5 hr. 18F present in the samples of 13N0;, radioactivity
of 18F/13N < 0.01 at the time of scanning, had no effect on the
results of the experiments, because the radioactivity of the
substances detected on the thin layer plates subsequently decayed
to background with the same half-life as 13N. During

electrophoresis, ammonium and nitrate migrate off the plate at

the negative and positive end, respectively (31).

 

RESULTS

+
Assimilation of 13NH4__

 

The organic products observed after various periods of
assimilation of 13NH;. are shown in Figure l and Figure 2a for
cells grown on nitrate, and in Figures 2b and c for cells grown
on urea or ammonium succinate as the sole nitrogen source.
After 10 s of assimilation, [13N]glutamine accounted for 96% of
the total radioactivity found in organic products soluble in 80%
methanol, for the nitrate-grown cells, and - similarly - for
93% of that total for cells grown with urea or ammonium
succinate. With longer periods of assimilation the percentage
of radioactivity in glutamine decreased, with a concomitant
increase in the percentage in other amino acids. Incorporation
of 13N into glutamine and glutamate was approximately linear
with time: after 10—15 min of assimilation (Table I), the
fractions of organic 13N in glutamine and glutamate were
approximately 70% and 20% for cells grown with any of the
nitrogen sources. Although the results varied quantitatively
from experiment to experiment, the amount of 13N in glutamine

was always higher than that in glutamate. A small percentage

of organic 13M was found in alanine and in an unidentified

144

145

. . . . l3 .
Figure 1. Scan of radioactiVity from N in electrophoreto-
grams of organic compounds extracted from
nitrate—grown, cultured tobacco cells with 80%

methanol after (a) 600, (b) 120, (c) 30, and

(d) 10 s of assimilation of 13NH41
13 + .
The same sample of NH was used for all four periods

4
of assimilation. The extracts, supplemented with ten

stable amino acids as markers, were applied to cellulosic
thin layer plates in a thin strip 8.5 cm from the

negative end (at the left) of the plate. Lipids were
displaced from the origin by chromatography in chloroform/
methanol (3/1, v/v). The extracts were then subjected

to electrophoresis at 3000 V for 11 min in 70 mM sodium
borate buffer, pH 9.2. The plates were scanned at 2 cm/min
(T = 3 s) from - to +, and were then dried and sprayed with
a solution of ninhydrin to localize the marker amino acids.
The 13N-containing substances in c and d co-migrated with
glutamine (at 9.6 cm) and glutamate (at 14.5 cm) during
electrophoresis, as did the two major 13N-containing
substances in a and b. The extracts in a and b also
contained 13N-labeled substances which migrated with alanine
(at 7.7 cm) and (appearing as a shoulder at the righthand
side of the glutamate peak) aspartate. A quantitative
evlauation of electrophoretograms such as those of Fig. l

is presented in Fig. 2.

146

I: N10. x Emu

I IONIC. X 2&0

1.. N10. x 2&0

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

BtO‘

OCS

LkCIKUQO‘Q‘k ‘Mx

6 3 .0 6 3 0 9 6 3 0 O
_ . a _ _ .u _ _ . M _ 2

a w. w C A. v

on M
.. .- .a
‘\\. I\‘.\ w

-.-- 4...! 1
.. :4: .- -...--. s r s

ooooooooooooooooo J IIIIIIII A

"..l' I '
\
V

,. . 1
II\\ II\\ lrl L 1' I\\\ I 0
fl fl flI "HI I
:I 1' #5

._

a _ _ F _ _ ¥

I O 2 I O 2 I 02 .—l 00
InIO_x 2&0 Inigx Emu Into.x Emu In10.x 2&9

DISTANCE (Cm)

147

Figure 2. Distribution of 13N in organic substances
soluble in 80% methanol after assimilation of
13NH;' for 10, 30, 120 and 600 s by cultured
tobacco cells grown on (a) nitrate, (b) urea
or (c) ammonium succinate as the sole nitrogen

source 0

Methanolic extracts were subjected to electrophoresis
at pH 9.2, the thin layer plates were scanned, and each
peak of radioactivity was integrated. A peak by peak
normalization was performed to correct for decay of 13N.
Values presented are the fraction of organic 13N migrating
during electrophoresis with glutamine (o), glutamate (c3)

and other organic compounds (A).

148

 

 

 

 

 

6
TIME(min)

 

 

 

 

 

 

 

 

 

_ _
0| 5 0| 5. 0

Zn. uo ZOE-04m...

 

149

.N can H mousmwh mo mocmmwa opp cfi omnfiuommo mm mommmooum mums mwamfimmo

 

 

 

m + ha m + mu m mauoa mumcHUUSm financesm
m H ma 3 H E o 3 sons
m H mm a H mo m S munnuaz
mumfimusHU wcwemusao mumxo GOMUMHflEflmm< szOHU now
use 2mH monomuuxo mo ucwoumm mo umnssz ca: monaom comouufiz
-

wcHE ma no 0H Mom .+m2ma mo coflumHHEwmwm nouns mumsmusHm mam ocwfimusHm cfi ZMH mo :ofiuomum com: .H anme

150

compound which migrated between arginine and alanine.

. . . . . . + . . . .
Inhibition of ASSimilation of 13NH, by Methionine Sulfox1mine

'3

 

and Azaserine

 

When 1 mM methionine sulfoximine (MSX) was added to a
suspension of urea-grown cells 15 to 20 min before it was exposed
to 13NH;: formation of 13N-labeled glutamine and glutamate was
decreased by 70% and 29%, respectively, in one experiment, and
by 99% and 88%, respectively, in another. When nitrate-grown
cells assimilated 13NHZ' in the presence of 1 mM MSX, the amount
of 13N in glutamine after 10 min of assimilation was decreased
by 95% whereas the amount of 13N in glutamate was decreased by
82% (Table II). Similar results were obtained when the experi-
ment was repeated. When cells grown with ammonium succinate were
treated with MSX at that same concentration, inhibition of 82%
and 28% for the first experiment and 49% and 15% for the second
experiment for glutamine and glutamate, respectively, were
obtained. Although there was considerable variation in the
extent of inhibition by MSX of assimilation of ammonium by cells
grown with urea or ammonium succinate, the percentage decrease
of labeling of glutamine was in all cases (including cells
grown with nitrate) greater than the percentage decrease of
labeling of glutamate. When 1mM azaserine was added to a
suspension of nitrate-grown cells 15 to 20 min before exposure

13

to NHJ, the formation of [13nglutamate was inhibited by 99%

while the formation of [13N]glutamine was inhibited by 56%.

 

151

Table II. Effect of 1 mM methionine sulfoximine on the
13 . .
appearance of N in glutamine and glutamate in

. +
suspenSions of tobacco cells exposed to 13NH and

 

 

 

 

4
13 -a
N03.
Nitrogen Source Source Percent inhibition of
13 -
for Growth of 13N appearance of N in
Glutamine Glutamate
Nitrate 13NHZI
Exp. 1 95 82
Exp. 2 94 83
Nitrate 13Nog'
Exp. 3 82 99
Exp. 4 89 93
+
Urea 13NH4
Exp. 1 79 29 I
Exp. 2 99 88
. . 13 +
Ammonium succ1nate NH4
Exp. 1 82 28
Exp. 2 49 15
aFor each experiment, 13N was assimilated simultaneously +

 

methionine sulfoximine for 10 min and the amounts of 13N_in
glutamine and glutamate determined as described in the legends
of Figures 1 and 2.

152

Similar results were obtained when the experiment was repeated

(Table III).

Pulse-Chase Experiments

 

A pulse-chase experiment was performed by exposing a

. . 13 + .
suspenSion of nitrate-grown cells to NH4 for 1 min, whereupon

a solution of NH4C1 was added to obtain a final concentration

of 10 mM NHJZ The cells were extracted 10, 30, and 60 5 later.
After the l-min chase, the percent of total organic 13N in
glutamine decreased, whereas that in glutamate plus alanine
increased, by 21% (Fig. 3). In a similar experiment, in which
the chase lasted 2 min, the decrease in [13N]glutamine and the
increase in [13nglutamate plus [13N]alanine were both 26% of
the total organic 13N. Chases longer than 2 min did not result

in additional loss of 13N from glutamine or additional increase

of 13N in glutamate or alanine.

Assimilation of 13N0;;_

 

The organic products observed after various periods of
assimilation of 13N05- by cells grown on nitrate are shown in
Figure 4. Usually, 13N did not appear in glutamate until 1 min,
although traces of glutamine and of an unknown compound (y)
which migrates between alanine and arginine during electro-
phoresis at pH 9.2 did appear before this time. At 1 min, most

of the 13N was found in glutamine (75%) and much less was in

glutamate (15%). After longer periods of assimilation, the

153

Effect of 1 mM azaserine on the appearance of 13N in

 

 

Table III.
glutamine and glutamate in suspensions of nitrate-
grown tobacco cells exposed to 13NHZ'.a
. . . . 13 .
Percent inhibition of appearance of N in
Glutamine Glutamate
Exp. 1 56 99
Exp. 2 47 91

 

aExperiments were performed simultaneously :_azaserine for 10 min,
and the amount of 13N in glutamine and glutamate determined as
described in the legends of Figures 1 and 2.

 

154

Figure 3. Distribution of 13N in organic substances
extracted with 80% methanol from nitrate-grown,
cultured tobacco cells when assimilation of
13NH2. for 1 min without supplemental NH;-
followed by assimilation in the presence of

+
10 mM NH4 for 0, 10, 30 and 60 s.

was

The fractions of organic l3N migrating with glutamine
(o), glutamate (c3) and alanine (A) during electrophoresis

are presented, evaluated as for Fig. 2.

155

 

FRACTION OF ”N

in

 

 

 

 

 

mo
OIDwm pmaooa my

 

156

. . . . 13 .
Figure 4. Distribution of N in organic substances soluble

in 80% methanol after assimilation of 13N05- for
l, 2, 6, and 15 min by cultured tobacco cells

grown on nitrate as the sole nitrogen source.

The fractions of organic 13N migrating during
electrophoresis with glutamine (o), glutamate (t3) and
other organic compounds (A) are presented, evaluated as

for Fig. 2.

157

 

 

 

 

J

‘I

 

 

 

N

2|

'9.
:IO NOIIOVHJ

l5

l0

TIME (min)

 

158

ratio of 13N in glutamate to 13N in glutamine was higher. With
the exception of the initial lag in formation of [13N]glutamate,
incorporation of 13N into both compounds was approximately linear
with time. An average of values taken from 23 experiments
performed for 6 to 15 min showed that 45 :_15% of the organic

3
N extracted was present in glutamate whereas 39 :_l4% of the

13 . . . 3
extracted N was present in glutamine. The remainder of the 1 N

was usually divided equally between compound y and alanine.

Addition of 1 mM nog' did not stimulate incorporation of lBNOéZ

The average fraction of the 13N supplied which appeared in

organic products after 10 to 15 min of assimilation was 4.5% when

4. .-
13NH4 was used, but only 0.3% when 13N03 was used, corrected

for decay. A sample of 13N03 was purified with the HPLC by

injecting the entire sample on the column and collecting only the

13N0;' fraction. When an assimilation experiment was performed

with the HPLC-purified sample, the appearance of 13N in organic

products was similar to that shown in Figure 4, for which 13N0;'

had been purified by the standard method.

When nitrate-grown cells were exposed to 13N0;' for 10 min

in the presence of 1 mM methionine sulfoximine, incorporation of
13M into glutamine and glutamate was decreased by an average
of 86% and 96%, respectively, in two experiments (Table II).

A pulse-chase experiment was performed by exposing a cell

was
3

suspension to 13N0;' for 5 min, after which stable N0

added to obtain a concentration of 10 mM. Assimilation con-

3 for 2 and 5 min, and was

tinued in the presence of 10 mM NO

 

159

then stopped by extraction of the cells. During the 5-min
chase, the percent of the total organic 13N in glutamine
decreased, and that in glutamate plus other amino acids

increased, by 19% (Fig. 5).

160

. . . . 13 . .
Figure 5. Distribution of N in organic substances
extracted with 80% methanol from nitrate-grown,

cultured tobacco cells when assimilation of

13N0;- for 5 min was followed by assimilation

in the presence of 10 mM N03

for 0, 2, and 5

min.

The fractions of organic 13N migrating during
electrophoresis with glutamine (o), glutamate (c3) and

other organic compounds (A) are presented, evaluated as

for Fig. 2.

FRACTION 0F'3 N

161

 

 

 

 

 

 

2
CHASE PERIOD (min)

I I
<\
.5-
_.o
a
t/
A- 1 1G
0L 1 1
5

 

 

DISCUSSION

The first major product of assimilation of 13NH2' by

tobacco cells, previously grown on nitrate, urea, or ammonium
succinate as the sole source of nitrogen, is glutamine. Gluta-
mate is the next most highly l3N-labeled amino acid, followed by
alanine. Although 13N in glutamine, as a percent of the total
extracted organic 13N, decreases with increasing periods of
labeling, the amount of 13N in glutamine is always higher than
the amount in glutamate. This pattern of labeling of amino
acids is similar to that found in roots of rice seedlings fed

15

+
NH4 for periods of 5 min and longer (32).

Methionine sulfoximine, a specific inhibitor of glutamine
. . . . . . . +
synthetase in vttro (24), inhibits the incorporation of 13NH4

into glutamine more extensively than it inhibits the

incorporation of 13NH2' into glutamate, regardless of whether the

- +
3, NH or urea (Table II). If there were

cells were grown on NO 4

no glutamate dehydrogenase activity one would expect the
inhibition by MSX of incorporation into glutamate to be at

least as great as the inhibition of incorporation into glutamine.
The data in Table II thus suggest that a fraction - in the

case of the nitrate-grown cells, a small fraction - of the

162

 

163

glutamate formed is synthesized by glutamate dehydrogenase.
However, an alternate explanation would be that a portion of

the [13nglutamine formed from 13NH;, but not from 13N0;(see
below) is used preferentially for glutamate synthesis. MSX,

at concentrations of 1 mM or 0.1 mM, is completely inhibitory

to growth of the cells on all three nitrogen sources (unpublished
results). A recent paper which studied the effect of MSX on

15 + . . . . . .
NH ass1milation in rice seedling roots (1) reported results

4
similar to ours. There was a 90% inhibition of incorporation
of 15N from ISNHZ' into glutamine when the seedlings were
pretreated with 5 mM MSX for 10 min and fed ISNHJ- for 1 hr.
Incorporation of 15N into glutamate was inhibited by 77%.
Azaserine, which inhibits all glutamine amide transfer reactions
including that catalyzed by glutamate synthase (20), inhibits
glutamate synthesis extensively in cells grown on nitrate (Table
III). This observation implies, as do the results with MSX
(see above), that in nitrate-grown cells, most of the glutamate
is formed at the expense of glutamine. An alternate
explanation for why there is very little incorporation of 13N
into glutamate in the presence of azaserine could be that
glutamine synthetase, because of its lower Km for ammonium, is
successfully competing with glutamate dehydrogenase and is
incorporating most of the label into glutamine.

When lBNHZ' is assimilated for l min and then chased with

13 . .
10 mM NHJZ only 21 to 26% of the. N in glutamine is

transferred into glutamate and alanine. Several possible

164

explanations as to why there is no further decrease in label in
glutamine with increasing time of chase are: that the glutamine
pool is expanding during the chase, so that the amount of [13N]-
glutamine metabolized decreases with time; that radioactivity
chased from glutamine (amide) to glutamate is being chased back
into glutamine (a-amino); or that much of the glutamine may have
moved into a metabolically inactive pool. Perhaps after shorter
periods of pulse-labeling, not technically feasible in these
experiments, a greater fraction of the 13N in glutamine would
have been transferred to glutamate during a chase. One would
expect that if glutamine and glutamate were formed independently,
3N would be lost in parallel from these two amino acids during
a chase. Thus, the fact that radioactivity in glutamate
increases relative to that in glutamine during the chase is
consistent with operation of the glutamine synthetase-glutamate
synthase pathway. Yoneyama and Kumazawa (32) transferred roots
of rice seedlings from medium containing 15NH+ to medium.with

4

unlabeled NHZZ In these experiments a 2-h pulse was followed

by chases of 30-min duration. Although 15N in glutamine
decreased, a concomitant increase in [lsN]glutamate was not
observed.
. . . . 13 -
The first major product of aSSimilation of NO in cells

3
. . . . 13 -+
grown on nitrate as the sole nitrogen source is, as with NH4,

[13N]glutamine. Although incorporation of 13H into glutamine
was observed after the shortest periods of assimilation studied

(30 s), it usually takes 1 min for label in glutamate to become

165

measurable. At all times of assimilation, a larger fraction of
13 . . . . 13 + .
the N supplied appears in amino aCids when NH4 13 used as

the nitrogen source than when 13nog' is used. Yoneyama and
Kumazawa (32,33) also observed considerably higher incorporation
15 . . . . .
of N into organic products in roots of rice seedlings when
15

. . + . -
these were supplied With NH rather than with 15NO

4 3. It has

been reported that the tobacco cells used in our experiments
(and possibly higher plants in general) contain both metabolic
and storage pools of nitrate (8). The lower assimilation rate
for nitrate may be attributable to the transport of a large
fraction of the nitrate to the storage pool. Alternatively, the
rate of permeation or of reduction of nitrate may limit the rate
of its assimilation.

After assimilation for 10 min or longer, the ratio of
[13nglutamate to [13nglutamine is higher when the tobacco
cells are given l3nog' as the nitrogen source than when 13NHZ'
is the nitrogen source. The greater relative incorporation of
13N into glutamate is more extensively inhibited than is
incorporation into glutamine (Table II). These results indicate
that assimilation of NH2- derived from nitrate proceeds
essentially completely via the glutamine synthetase-glutamate

synthase pathway. The apparently lesser contribution of

o - . +
glutamate dehydrogenase to assimilation of 13NO -derived 13NH4

3
. 3 + .
than to assimilation of exogenously applied 1 NH4 may pOSSibly
13 +
reflect the buildup of a lower intracellular pool of NH4 in

the former case than the latter; the activity of glutamine

 

166

synthetase and glutamate synthase would in that former case be
favored over activity of glutamate dehydrogenase because of the

higher Km of glutamate dehydrogenase for ammonium.

The results of the chase experiment with 13NO-' are

3
. . . . 13 +
Similar to the results of the chase experiment with NH4,

and also favor the glutamine synthetase-glutamate synthase

pathway. When 10 mM N0;' is used to dilute the 13N0;' after a

5-min assimilation period, a small decrease (19%) in the percent
of the total extracted 13N in glutamine and a corresponding
increase in the percent of the 13N in glutamate are detected
after 5 additional min. Yoneyama and Kumazawa (33) also

observed an increase in [lleglutamate together with a decrease

in [lsN]glutamine, when roots of rice seedlings which had

assimilated lsNog' for 2 h were transferred to Nogz

 

BIBLIOGRAPHY

10.

11.

12.

13.

14.

15.

16.

BIBLIOGRAPHY

Arima, Y., Kumazawa, K., Plant Cell Physiol., 88, 1121-1126
(1977).

Austin, S. M., Galonsky, A., Bortins, J., Wolk, C. P., Nucl.
Inst. Meth., 126, 373-379 (1975).

Bauer, A., Urquhart, A. A., Joy, K. W., Plant Physiol., 88,
915-919 (1977).

Carangal, A. R., Varner, J. E., Plant Physiol., 88, (Suppl.h
xi (1959).

Dougall, D. K., Biochem. Biophys. Res. Commun., 88, 639-646
(1974).

Dougall, D. K., Bloch, J., Can. J. Bot., 88, 2924-2927(19761
Elliott, W. H., J. Biol. Chem., 201, 661-672 (1953).

Ferrari, T. E., Yoder, O. C., Filner, P., Plant Physiol.,
88, 423-431 (1973).

Filner, P., Exp. Cell Res., 88, 33-39 (1965).
Filner, P., Biochim. Biophys. Acta, 118, 299-310 (1966).

Gersberg, R., Krohn, K., Peek, N., Goldman, C. R., Science,
192, 1229-1231 (1976).

Givan, C. V., Planta, 122, 281-291 (1975).
Haystead, A., Planta, 111, 271-274 (1973).

Heimer, Y. M., Filner, P., Biochim. Biophys. Acta, 230,
362-372 (1971).

Lea, P. J., Miflin, B. J., Nature, 251, 614-616 (1974).

Lewis, 0. a. M., J. Exp. Bot., 39, 361-366 (1975).

167

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

168

Lewis, 0. A. M., Berry, M. J., Planta, 125, 77-80 (1975).

Lewis, 0. A. M., Pate, J. S., J. Exp. Bot., 88, 596-606
(1973).

Meeks, J. C., WOlk, C. P., Thomas, J., Lockau, W., Shaffer,
P., Austin, 8. M., Chien, W-S., Galonsky, A., J. Biol.
Chem., 252, 7894-7900 (1977).

Miflin, B. J., Lea, P. J., Biochem J., 149, 403-409 (1975).
Miflin, B. J., Lea, P. J., Phytochem.,-88, 873-885 (1976).
O'Neal, D., Joy, K. W., Nature New Biol., 246, 61-62 (1973).

O'Neal, D., Joy, K. W., Arch. Biochem. Biophys., 159, 113-
122 (1973).

O'Neal, 0., Joy, K. w., Plant Physiol., 21: 773-779 (1974).

Thomas, J., Wolk, C. P., Shaffer, P. W., Austin, S. M.,
Galonsky, A., Biochem.Biophys. Res. Commun., 82, 501-507
(1975).

Thomas, J., Meeks, J. C., Wolk, C. P., Shaffer, P. W.,
Austin, S. M., Chien, W-S., J. Bacteriol., 129, 1545-1555
(1977). “"

Tiedje, J. M., Smith, M. S., Firestone, M. K., Betlach, M.
R., Firestone, R. B., in Proc. Int. Symp, Microbiol. Ecology
(Dunedin, New Zealand), Springer Verlag, Berlin, In press
(1979).

Tilbury, R. S., Dahl, J. R., Radiation Research Society 25th
Annual Meeting, San Juan, Puerto Rico, Paper D(h)-7, (1977).

Webster, G. C., Varner, J. E., Plant Physiol., 88, 724-727
(1953).

W61k, C. P., Austin, S. M., Bortins, J., Galonsky, A.,
J. Cell Bio., 88, 440-453 (1974)-

Wblk, C. P., Thomas, J., Shaffer, P. W., Austin, S. M.,
Galonsky, A., J. Biol. Chem., 251, 5027-5034 (1976).

Yoneyama, T., Kumazawa, K., Plant Cell Physiol., 88, 655-661
(1974).

Yoneyama, T., Kumazawa, K., Plant Cell Physiol., 191,21'25
(1975).

169

34. Campbell, N. E. R., Dular, R., Lees, H., Can. J. Microbiol.,
88, 587-599 (1967).

35. Nicholas, D. J. D., Silvester, D. J., Fowler, J. F., Nature,
189, 634-636 (1961).

36. Ogg, R. A., J. Chem. Phys., 88, 613 (1947).

37. Ruben, S., Hassid, W. Z., Kamen, M. D., Science, 88, 578-
579 (1940).