Citation
Adaptation of membrane-bound enzymes to ethanol

Material Information

Title:
Adaptation of membrane-bound enzymes to ethanol
Creator:
Dickens, Benjamin Fisher, 1951- ( Dissertant )
Ingram, Lonnie O. ( Thesis advisor )
Hoffman, Edward M. ( Reviewer )
Allen, Charles M. ( Reviewer )
Sisler, H. H. ( Degree grantor )
Place of Publication:
Gainesville, Fla.
Publisher:
University of Florida
Publication Date:
Copyright Date:
1978
Language:
English
Physical Description:
vii, 58 leaves : ill. ; 28 cm.

Subjects

Subjects / Keywords:
Adenosine triphosphatases ( jstor )
Alcohols ( jstor )
Enzymes ( jstor )
Ethanol ( jstor )
Fatty acids ( jstor )
In vitro fertilization ( jstor )
Incubation ( jstor )
Lipids ( jstor )
Membrane transport proteins ( jstor )
Sodium ( jstor )
Alcohol -- Physiological effect ( lcsh )
Dissertations, Academic -- Microbiology and Cell Science -- UF
Lipid membranes ( lcsh )
Microbiology and Cell Science thesis Ph. D
Genre:
bibliography ( marcgt )
non-fiction ( marcgt )

Notes

Abstract:
Previous studies have shown that the proportion of unsaturated fatty acids increases in _E^he£ichia_coli_during growth in the presence of ethanol. These lipid changes presumably represent an adaptive response to compensate for the direct physical effect of ethanol on the membrane lipid bilayer. The effects of growth in the presence of ethanol on two membrane-bound enzymes, the lactose permease and adenosine triphosphatase, were investigated. All but one of the properties of these membrane-bound enzymes, the activity of membranebound ATPase with Ca as a cofactor, were found to adapt to the presences of ethanol. The role of new protein synthesis and the ethanol-induced changes in the bulk lipid composition in the adaptation of membrane-bound enzymes to ethanol was also investigated, Results of these investigations led to a model proposing that ethanol's effect on both membrane-bound enzymes and the adaptation of these enzymes to ethanol is mediated through the enzymes annular lipids.
Thesis:
Thesis--University of Florida.
Bibliography:
Bibliography: leaves 54-57.
General Note:
Typescript.
General Note:
Vita.
Statement of Responsibility:
by Benjamin Fisher Dickens, Jr.

Record Information

Source Institution:
University of Florida
Holding Location:
University of Florida
Rights Management:
Copyright [name of dissertation author]. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
Resource Identifier:
000081135 ( alephbibnum )
05086234 ( oclc )
AAJ6453 ( notis )

Downloads

This item has the following downloads:


Full Text










ADAPTATION OF MEMBRANE-BOUND
ENZYMES TO ETHANOL









By

BENJAMIN FISHER DICKENS, JR.


A DISSERTATION PRESENTED TO THE GRADUATE COUNCIL OF
THE UNIVERSITY OF FLORIDA
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE
DEGREE OF DOCTOR OF PHILOSOPHYY


UNIVERSITY OF FLORIDA














DEDICATION


To my wife, Bonnie, who stood by me through the years it

took to achieve this goal; to my son, Brian, who has brightened

my life; to my parents, to whom I owe more than I will ever be

able to repay; this dissertation is dedicated to all of you.














ACKNOWLEDGMENTS

I would like to offer special thanks to Dr. L. O. Ingram, who

as my major professor offered me the benefit of his knowledge and

experience. I would also like to express my appreciation to Dr. E.

M. Hoffmann and Dr. C. M. Allen Jr. for serving on my graduate

committee and for their assistance during my graduate career. In

addition, I am very grateful to the faculty of the Department of

Microbiology and Cell Science who were always willing to share

their time and resources.














TABLE OF CONTENTS


ACKNOWLEDGMENTS .......................... iii

KEY TO ABBREVIATIONS ......................... v

ABSTRACT ................................... vi

INTRODUCTION ............................... 1

MATERIAL AND METHODS ...................... 3
Lactose Permease .......................... 3
ATPase .................................. 4
Arrhenius Plots of Lac Permease and ATPase .... 5
Fatty Acid Analysis ......................... 6
Chemicals ................................ 6

RESULTS ....................................... 8
Effect of Ethanol on the Lac Permease ......... 8
Effect of Ethanol on ATPase ................. 11
Arrhenius Plots ............................ 22
Reconstitution of Membrane-bound ATPase ..... 33
Cooperativity of ATPase Inhibition by Sodium ... 36
In Vitro Acclimation of ATPase ............... 46

DISCUSSION ................................. 49

BIBLIOGRAPHY .................................. 54

BIOGRAPHICAL SKETCH .......................... 58














KEY TO ABBREVIATIONS


ATP: Adenosine Triphosphate

ATPase: Adenosine Triphosphatase

Ea: Energy of Activation

lac permease: Lactose Permease

ONPG: o-nitrophenol ,-D-galactopyranoside

S. D.: Standard Deviation

16:0 : Palmitic acid

18:1 : Oleic acid














Abstract of Dissertation Presented to the Graduate Council
of the University of Florida in Partial Fullfilment of the Requirements
for the Degree of Doctor of Philosophy



ADAPTATION OF MEMBRANE-BOUND ENZYMES TO ETHANOL

By

BENJAMIN FISHER DICKENS JR.





Chairman: Lonnie O. Ingram
Major Department: Microbiology and Cell Science

Previous studies have shown that the proportion of unsaturated

fatty acids increases in Escherichia coli during growth in the presence

of ethanol. These lipid changes presumably represent an adaptive

response to compensate for the direct physical effect of ethanol on

the membrane lipid bilayer. The effects of growth in the presence of

ethanol on two membrane-bound enzymes, the lactose permease and

adenosine triphosphatase, were investigated. All but one of the prop-

erties of these membrane-bound enzymes, the activity of membrane-

bound ATPase with Ca++ as a cofactor, were found to adapt to the

presence of ethanol. The role of new protein synthesis and

the ethanol-induced changes in the bulk lipid composition in the

adaptation of membrane-bound enzymes to ethanol was also investigated.








Results of these investigations led to a model proposing that ethanol's

effect on both membrane-bound enzymes and the adaptation of these

enzymes to ethanol is mediated through the enzymes annular lipids.















INTRODUCTION

Living organisms display a remarkable ability to adapt their

membranes to environmental changes such as temperature, by altering

their lipid composition (Fulco, 1974). These lipid changes are thought

to maintain the proper environment within the membrane for biological

function (Sinensky, 1974). Ethanol has been shown to directly inter-

calate within biological membranes altering membrane organization (Hill

and Bangham, 1975; Lee, 1977; and Seeman, 1972). Growth of E. coli

in the presence of ethanol results in the synthesis of membrane lipids

with an altered fatty acid composition, analogous to the changes observed

following a decrease in growth temperature (Ingram, 1976).

The enzymes of fatty acid synthesis appear to be a major site for

the regulation of lipid composition in E. coli (Cronan, 1975). Temperature

induced changes in the fatty acid composition are mediated in part through

direct effects on these soluble enzymes (Cronan, 1975). Investigation

of the mechanism of the ethanol-induced fatty acid changes indicate

that the site of action is also the soluble fatty acid synthetase enzyme

(Buttke and Ingram, 1978). This has recently been confirmed in our

laboratory in vivo.

Ethanol has been shown to affect the activity of a variety of

membrane-bound enzymes. Membrane-bound ATases in eukaryotic






2

cells have been frequently studied (Grisham and Barnett, 1972; Mitjavila

et al., 1976; Post et al., 1972; and Sun, 1976). Ethanol generally

inhibits the oubain-sensitive (Na,K)-ATPase but has little effect on

the activity of the Ca -ATPase. Transport systems have also been

studied and are usually inhibited by ethanol in eukaryotic cells (Chang

et al., 1967; Fox et al., 1978; Hoyumpa et al., 1977; Israel et al.,

1968; and Worthington et al., 1978). In E. coli, alcohols of different

chain lengths have opposite effects on the /-galactoside transport

enzyme, the lac permease. Alcohols of chain lengths greater than four

stimulate lac permease activity (Sullivan et al., 1974) while alcohols

of shorter chain lengths are inhibitory (Fried and Novick, 1973). A

similar differential effect of short and long chain alcohols has been

reported on fatty acid composition (Ingram, 1976).

The fatty acid changes induced by ethanol in E. coli have been

proposed as part of an adaptive response, compensating for the direct

physical effects of ethanol (Ingram, 1976). The similarities between

the lipid changes induced by ethanol to those resulting from a decrease

in growth temperature suggest that both may be involved in the mainte-

nance of the proper lipid environment for membrane function. In this

paper, we have investigated the significance of ethanol-induced

changes for membrane function in E. coli using two enzymes, ATPase

and lac permease. The effects of ethanol on these enzymes were

compared between cells grown in the presence and in the absence of

ethanol.













MATERIALS AND METHODS


Lactose Permease

Escherichia coli K12 strain Kl-221 (lac i ), generously donated by

Dr. R. P. Boyce (University of Florida, Gainesville, Florida, 32611),

was used to examine the effect of ethanol on the lac permease. Cultures

(250mL) were grown in mineral salts medium M63 (Miller, 1972) supple-

mented with succinic acid (5g/L), L-threonine (40mg/L), L-tyrosine

(20mg/L), L-proline (30mg/L), L-histidine (22mg/L), uracil (40mg/L)

and thiamine (20mg/L) in oversized culture tubes with continuous

aeration or in 2-L flask with continuous shaking at either 30 OC or 37 C.

Lac permease activity was assayed as described by Fried and Novick

(1973). Cells were harvested in exponential phase by centrifugation

and resuspended in inhibitor buffer. Portions of this cell suspension

were removed and supplemented with o-nitrophenol/- D-galactopyranoside

(ONPG). The reaction was stopped by the addition of calcium bicarbonate

and the o-nitrophenol concentration determined spectrophotometrically

at 420nm. Formaldehyde treated controls were used to correct for cryptic

transport (Kepes, 1971).

In contrast to the above assay, the in vivo permease activity was

determined in actively growing cells by supplementing ImL samples of

growing cultures with ONPG (1.85mM) and continuing to incubate under

growth conditions for 3-7 minutes. Pernease activity was measured as

3








o-nitrophenol produced using a formaldehyde control. Total

9-galactosidase activity was measured by the procedure of Putman

and Koch (1975). Cell numbers were estimated by turbimetric

measurements.

ATPase

Escherichia coli KI2 strain CSH-2, obtained from the Cold Spring

Harbor Laboratory (Cold Spring Harbor, N. Y.), and a derivative of

this strain blocked in fatty acid degradation (Buttke and Ingram, 1978),

strain TB-4 (fad E), were used to examine the effects of ethanol on

ATPase. Cultures (250mL) were grown in glucose-supplemented

Luria broth (Luria and Delbruck, 1943) in 2-L flask with continuous

agitation at either 30 C or 37 C. Cells were harvested in exponential

phase (2 x 108 cells/mL) by centrifugation and resuspended in a

minimum of buffer, lysed using a French pressure cell and the membranes

prepared as described by Futai et al. (1974). ATPase activity was

assayed as described by Evans (1969) with corrections for spontaneous

hydrolysis of ATP. The reaction was terminated by the addition of

trichloroacetic acid. Liberated inorganic phosphate was determined

colorimetrically by the method of Rathburn and Betlach (1969). Protein

was determined by the method of Lowry et al. (1951). The specific

activity of the ATPase preparations was 37.6 f5.6 micromoles ATP

hydrolysed/(mg protein x hour) at 300C.

Reconstitution experiments were performed with ATPase as described

by Rosen and Adler (1975). ATPase was solubilized using low ionic









strength buffers. This procedure solubilizes the FI ATPase with

bound annular lipids (Peter and Ahlers, 1975). Stripped membranes

were prepared by repeated extraction with the solubilizing buffer and

hydrolysed less than 0.20 micromoles of ATP/(mg protein x hour) at

300C. Reconstituted membranes used for Arrhenius plots hydrolysed

about 24 micromoles of ATP/(mg protein x hour) at 300C.

Arrhenius Plots of Lac Permease and ATPase

During the determination of permease activity, freshly prepared

suspensions of cells were held in inhibitor buffer at 0C. Incubations

were performed using individual water baths for a total of 10 to 12

temperature points per plot. During the determination of membrane-

bound ATPase activity, fresh membrane preparations were held at 00C.

During the determination of soluble ATPase activity, freshly solubilized

enzyme was held at room temperature. For ATPase, fifteen temperatures

were assayed concurrently in duplicate using a linear temperature

gradient block. Slopes were determined by least square regression

analysis. The intercepts were determined mathematically by setting

the equations describing each line (above and below the break) equal

at log activity and solving for temperature. Coefficients of correlation

were determined for each line and ranged from 0.985 to 0.998 for the

lac permease and 0.990 to 0.999 for ATPase activity. Omission of the

two points nearest the break had little effect on these slopes.

The temperature gradient block used for Arrhenius plots of ATPase

consists of an aluminum block 22.5 x 4 x 4 inches. An array of inch







6

holes (15 x 3) was centered in this block (2.5 inches deep) Pieces

(1.5 inches) were cut from each end, routed to form two water channels

and fitted with inlet and outlet connections. These were bolted back

on the block using a rubber gasket as a seal. A Haake E52 thermostated

circulator was connected to one end (hot water) and a Neslab RTE-3

circulator was connected to the other end (cold water/ethylene glycol).

The block was encased in 1.0 inch of styrofoam with corresponding

holes, mounted in a wooden frame and bolted on an Eberbach recipro-

cating shaker (Scientific Products, Orlando, Florida). The temperature

ranges in the block were established by adjusting the circulator tempera-

tures and the block formed a stable gradient within two hours. The

temperature gradient as measured in each well using a NBS standard

thermometer did not deviate from linearity with a coefficient of

correlation greater than 0.9999. Wells accommodate 13 x 50 mm tubes.

Approximately 0.5 mL of water was added to each well to ensure good

thermal contact between the block and the sample tube.

Fatty Acid Analysis

Lipids were extracted from cells which had been inactivated with

trichloroacetic acid (5%) as described by Kanfer and Kennedy (1963).

Fatty acids were transesterified and analysed by gas chromatography

(Ingram, 1976). Compositions are reported as percentage of peak area.

Chemicals

All bicchemicals used in this study were obtained from Sigma

Chemical Company (St. Louis, Missouri). During the course of this






7

study Cantley et al. (1977) found that the ATP from Sigma used in this

report contained a potent inhibitor of (Na,K)-ATPase. This inhibitor,

an inorganic vanadium compound, does not effect the Ca -ATPase

activity (Cantley et al., 1977) or bacterial ATPase. This was confirmed

in our laboratory using E. coli ATPase with the vanadium free ATP which

was also purchased from Sigma.











RESULTS

Effect of Ethanol on the Lac Permease

Ethanol causes a dose dependent inhibition of the lac permease

(figure 1). Cryptic transport (diffusion and other transport systems)

is slightly stimulated by ethanol. These effects are reversible upon

removal of ethanol by washing. A variety of other moderately hydro-

phobic compounds have been shown to induce changes in E. coli lipid

composition alalogous to ethanol (Ingram, 1977). Of these, acetone

(20g/L), dimethyl sulfoxide (74g/L), methanol (32g/L) and dioxane

(20g/L) were tested for their effect on the lac permease. In all cases,

these agents inhibited the permease activity, Sullivan et al. (1974)

have previously shown that long chain alcohols such as hexanol

stimulate permease activity. This stimulation by long chain alcohols

was confirmed in strain K1-221. Growth of E. coli in the presence of

these long chain alcohols results in changes in fatty acid composition

opposite to those caused by ethanol (Ingram, 1976). Chloroform (0.5g/L)

and amyl acetate (0.3g/L) induce changes in lipid composition like

hexanol (Ingram, 1977) and these agents also stimulate permease activity,

The initial inhibition of permease activity by ethanol is relieved

during subsequent growth in the presence of ethanol. Both the permease

and# -galactosidase from cells grown in the presence of ethanol exhibit






















Figure 1. Acute effect of ethanol on P-galactoside
uptake. Lac permease mediated transport, ; and cryptic
entry, O .

















800







600-

Or
bJx

00
.co



o j
0
OE
0

c O
200







1 2 3 5 7 9


ETOH (%)






11

the same specific activities as found in control cells. This suggests

that the recovery of permease activity does not simply result from the

synthesis of new protein. The time required for the recovery of

permease activity is directly related to alcohol concentration (figure 2A)

and inversely related to the growth rate (figure 3A). Changes in fatty

acid composition occur during this recovery period. The time required

for the completion of these changes was directly related to alcohol

concentration (figure 2B) and inversely related to growth rate (figure 3B).

In all cases, permease activity was restored prior to the completion of

these changes in bulk lipid composition. Thus the completion of these

changes in lipid composition is not essential for the recovery. Protein

synthesis in strain Kl-221 was inhibited by the addition of chloramphen-

ical (100mg/L) and by threonine starvation. Neither treatment prevented

the recovery of permease activity during incubation with ethanol,

although a longer time was required (figure 4). The decrease of the

permease activity with time in both the ethanol-treated and control

cells is presumably due to turnover of permease.

Effect of Ethanol on ATPase

The effect of ethanol on ATPase isolated from E. coli grown at 370 C

in the absence of ethanol is dramatically affected by the choice of

divalent metal cofactor (figure 5). With Mg ++ ethanol caused a dose-

dependent stimulation of activity. With Ca ++, ethanol caused the

opposite effect, a dose-dependent inhibition. This differential effect

suggests that ethanol is interacting in some way which alters the shape
























C0C

C ) 00'rU "
1- C
LoO
Cn

u C


u ( C


9 (a Q) _e ^ *^
CL 0
in g












o^ "
C-o C o Cj








> 0
0 o 0 m 0 1



(6 C CL







0 fi"0
Q E & r Oro
CCC>O 0 C)










tc" 3 38I
SC !O 0




iC C o C
















C;o a a So
C)0C



S- ^ o
0 -0
Iu 0
*5i- o~ - C
rC-.4) )0
2Z~dac
ZCC) C)~~ '
C) C h. N C)O Q
C2u~



--c Ch







13





0
'1*




0


E
uj


o


lii







NO ~ ~ I ON
en W












if I









LUj
o(DH






























G3Z1O0dG)H 9dNO























En In

ma -






iE 80 e
CL
o 6



30 m c -
C C < 0 o nt

- D 0- 1













~u u t
3 o nO ^
0 0 0a c


0j Z 0 0 0






.00 wE 2(D
,4 -C0 o T



0 0 0 -- *o & 0
C o 2






>-i 0 0 *
O r C 0 0< 0 0


00-, 00 .2 o
"ii0^







U tt 1C 0 i, 1
r0 d. '-i1E 001










0
O



co










J NOI ISOdVOO3

GIOV AIIVJ '4DS/lDsun

0
a1






E
0-

LiJ





-0




o o o a
r8o



(Jnoq x slI a801 /salowOuDU)

G3ZAKO1CAH 9dNO





























CS -Cg
.j4'>







-O 0-


a0)

















ac


m o
- 1





C0 0 0










co
O-U








z C-
I- U






























'-000" "
00.-





.C o 41 -
0 00 <
-- c I U






17
0
-Go



a,
m O





Co








I-
(d




o o
0 0





0


Lui


0o -





0 0

(Jqy ] WU/salIowouDu)

G 3 ZA"IOHG aH EdNO




















Figure 5. Effect of ethanol on membrane-bound ATPase
from cells grown at 37 C with Mg++ or Ca+ as a cofactor
assayed at 37 C. Control cells with Mg'+,*; control
cells with Ca ,A; ethanol grown cells with Mg+ ,0;
and ethanol grown cells with Ca++, .

















100








LJ 50
LU



Fr







1 I I I I
2 6 10
ETHANOL (%)






20

of the catalytic site of the enzyme. The Km of ATPase was found to be

unaltered by the inclusion of ethanol in the assay mixture.

Growth in the presence of ethanol results in changes in the lipid

composition which could also influence enzyme function (table I).

ATPase from ethanol-grown cells was equally sensitive to inhibition by

ethanol using Ca++ as a cofactor (figure 5). However, ATPase from

ethanol-grown cells was much less stimulated by ethanol using Mg++ as

a cofactor (figure 5). Thus changes have occurred during growth in the

presence of ethanol which partially ameliorate the effect of ethanol on

ATPase activity assayed with Mg

Growth of E. coli strain CSH-2 at 300 C results in a fatty acid

composition similar to that produced during growth at 370 C in the

presence of ethanol (table I). When ATPase from cells grown at 30 C

was assayed at 370 C using Mg++ as a cofactor, ethanol caused a

modest stimulation of activity, intermediate between cells grown

without ethanol at 37 C and 37 C ethanol-grown cells. This suggests

that fatty acid changes may be involved in ethanol resistance and a

possible relationship between ethanol and low temperature.

Ethanol has been shown to have little effect on the activity of the

Mg++ stimulated ATPase of mitochondria from rat kidney when assayed

at high temperatures (Hosein et al., 1977). This enzyme was inhibited,

however, at lower temperatures (Hosein et al., 1977). The ATPase of

E. coli with Mgg as a cofactor is stimulated at 37 C (growth temperature)

by ethanol, however a slight inhibition in activity by ethanol was


I




















C- C L . . Lo m 0 co


c. c o .j m .--
C* *o C-I CO CM CO * *



C- CO O C r C:O M cMt Col C )O rC



co c C C Cl C r) Cl O l O C


UO) C) m o c) m 3 CN C) C) CL

o lo 0 C c C) 0 cn c
(' U) C' C Co, C- Ic' C] C) C ]1

0C-]0.o -Clll C C <4'C' i I


u >






I C











El l
s C
Cl

.C C




<~ 4.


o C C=
C C C C C CC (.
C' C C' C C' C,-401
0 o~g o~5oo
SMSMM- H


co n coCoomo o m




- C-- C- C0 C' C-
C'] C'] ',
VS ca C co m"

u ()u c) E


co
T


C E
o
I-i


0
0

M0


0 0
r or
0 0(
g-z 14


CD



E

o
0
0






o
CO







S.
0





en
0








CT
10











0 -.




TC







22
observed when assayed at a lower temperature (figure 6). This inhibition

at a lower temperature (150 C) was relieved following growth in the

presence of ethanol.

Arrhenius Plots

Arrhenius plots are frequently used to study membrane-bound enzymes

(for review see Lenaz et al., 1975). Arrhenius plots of membrane-

bound enzymes typically display one or two discontinuities. Massey

et al. (1966) have shown a direct correlation between the break tempera-

ture in Arrhenius plots and a conformational change in a soluble enzyme.

The position of the break in membrane-bound enzymes is presumed to

also be due to a conformatioral change in the enzyme. Break temperatures

are dependent upon the specific protein and its bound annular lipids

(Hesketh et al., 1976). Only one break was observed in the Arrhenius

plots in this report. Based upon previous reports (Linden et al., 1973;

Morrisett et al., 1975; and Thilo et al., 1977) the second break would

be expected to occur at or below our lowest measured temperatures.

In cells grown at 42 C and in cells at 37 C with exogenous palmitic

acid (16:0) to boost the saturated fatty acid content, we observed a

second break with the ATPase (data not shown).

Arrhenius plots of lac permease have break temperatures which are

dependent upon the lipid composition of the membrane (Overath et al.,

1971). Cells grown at 300C contain a lower proportion of saturated

fatty acids than cells grown at 370 C (table I). As expected, the break




















Figure 6. Effect of ethanol on membrane-bound ATPase
from cells grown at 37 C with Mg++ as a cofactor and
assayed at 15C. Control cells,6; ethanol-grown cells, 0.



















70


30-


2 6 (
ETHANOL (%)


110 -







25

temperature of the 300 C-grown cells is lower than that for 370 C-grown

cells (figure 7A & B). Inclusion of ethanol in the assay mixture resulted

in an increase in the break temperature of these Arrhenius plots (figure

7A & B). This increase is similar to the effect caused by an increase

in the proportion of saturated fatty acids. The effects of ethanol on

Arrhenius plots of lac permease are summarized in table II. From these

results it appears that ethanol in some way stabilizes the form of the

lac permease with the higher energy of activation, increasing the tempera-

ture necessary for the conformation change to the form with the lower

energy of activation. Unlike the break temperature, however, the

energy of activation of the lac permease appears unaffected by the

addition of ethanol with one possible exception (table II), the lower Ea

of cells grown at 370 C in the absence of ethanol.

Growth in the presence of ethanol results in changes within the

membrane which prevent the ethanol-induced increase in break tempera-

ture (figure 7C). Arrhenius plots of ethanol-grown cells assayed in

the absence of ethanol display the same break temperature as control

cells assayed in the absence of ethanol, despite differences in fatty

acid composition (table I). The lipid composition of strain KI-221

grown at 30 oC without ethanol and ethanol-grown cells at 37 OC is very

similar (table I). However, the break temperature in these 30 OC-cells

is still increased by the inclusion of ethanol. These results suggest

that bulk lipid composition alone does not determine break temperature

or ethanol resistance.















































cn
~rfl

























-0~


U2 -

-, )
C



Cl)
:5
ro
*0
m010
.- C.)
















0
0

0 0


o o 0
x






I I





O










0 0 0 0
0 00




0 O
O










(inoq x Sllax O/SalOLUOUoU)
30 0 0o 0NO
0 I) 0
(JnoJ x !I~ OI/OIOWUDU
a *
av~.foa r OdN










28



S+1
I 4 CI o i U)LOc


M C C 0






o
i++1-1+1 +1-I+-1+1








< 0 01mm) -m
i-;- CIM C) fn CM

IN -
\ ac r oi c Un m ^ O o n a












I o)





)+1 ^+ +1 +1+1 I r

















2a1
>
M
ra I "S
































< S -^ T






29

The break temperature in Arrhenius plots of ATPase is less affected

by changes in bulk membrane lipid composition than lac permease

(figure 8) although lipids are required for ATPase activity (Peter and

Ahlers, 1974). As with the lac permease, the inclusion of ethanol in

the assay mixture results in an increase in the break temperature of

membrane-bound ATPase (figure 8A & B). ATPase from ethanol-grown

cells exhibits no such increase in break temperature when assayed in

the presence of ethanol (figure 8C). The lipid composition of strain

CSH-2 grown at 300C without ethanol and ethanol-grown cells at 370C

are very similar (table I). However, the break temperature in these

300C-cells is still increased by the addition of ethanol, suggesting

that changes in bulk lipid composition per se do not provide protection

against ethanol. The effect of ethanol on Arrhenius plot characteristics

of ATPase are summarized in table III. As shown in this table, ethanol

did not alter the energies of activation below the break temperature in

ethanol-grown cells. The similarities between the effect of ethanol on

the break temperature of the lac permease and that of membrane-bound

ATPase may reflect a general effect of ethanol on lipid/protein inter-

actions.

ATPase can easily be solubilized by treatment with low ionic

strength buffers (Futai et al., 1974). Arrhenius plots of solubilized

ATPase exhibit the same break temperature as the membrane-bound form

(table III). Similar results were found by Sinerez et al. (1973). Inclu-

sion of ethanol in the assay mixture caused an increase in the break



























-a
u~-


la







S0 m
to z
CUd

(a


H0 0.
-a

0
at- a



G u)~
F-.a

0C0





0

0d -0
a 0
0 Cd







ILiI
0 *,
2~04
Cd )~
Cd'~ u '-4
.- - 4-
t-W0
















U












o 0 0 n
o d 2 N i
N ,
o o o o 9 I



l I d ~ I


0
I o









0










0
i








q


0 0 0 0
0 0 0

(Jnoq x ulaioJd 5wu/saIOLUoj3LUj)

03ZKA-IOOAH dlV











O.0m 00 om N c NO N m c o

C- cNi 1 oC m r Coo-C
+ +1+ 1+1 + +1+41+1 + +1+1+1 I+1 +1+1+1+1 +1+1+1+1 I
( -c r Io n m 0 0 o3 3 0 m C
O U m L c'Jo"uooa enC N L0Lo
-)--+i ) ecnocN en cM cmeN cl c ci in nJ m mcc


I 1 c O ujjC' O CC C C Omu n Cooc
L- I C) M00 OOm(00 0aO)MW CMn ma)C 0mo00M

+ + I +I1+1+ +1+1+1+ +1+1+1+1 +II+ II+1 +1+l+1+1
S L .( CCM- e C0 C ] O C- ,-
Co 0 (0o 0 o co co o a) 0 o o 0 -i



II





)D D 000 0 000N- 0000 0000 0000 CD

- 1 : + : + ++ +14 +1+1 +1+ 1+1+ 1 + +I +1+1+ 4


C -





0 0 0 0 0 0 0

cco do o(oo oo
+ +D








r5r
---a 0 0 0 0 0 0 0 + OI







I -













a CI



E-O -OM C








(D 00
CC 0 <






33

temperature of soluble ATPase (table III) analogous to that observed

with membrane-bound ATPase. This indicates a direct effect of ethanol

on the soluble ATPase complex. Thus the effect of ethanol on Arrhenius

plots is not mediated through the interaction of ethanol with bulk

membrane lipids. Soluble ATPase isolated from ethanol-grown cells is

also resistant to the effect of ethanol (table III), indicating that bulk

membrane is not directly responsible for the protection against ethanol.

E. coli strain WN I fails to alter its lipid composition when grown

at different temperatures (Gelmann and Cronan, 1972) or in the presence

of ethanol (table I). The membrane-bound ATPase from this organism

displays a break temperature in its Arrhenius plot which is increased

by the inclusion of ethanol (figure 9A). The break temperature from

cells grown in the presence of ethanol is unaffected by ethanol (figure

9B) despite the failure of this strain to alter its bulk fatty acid composi-

tion. Again, changes in bulk lipid composition do not appear essential

for the amelioration of the ethanol effect on Arrhenius plots of ATPase.

Reconstitution of Membrane-bound ATPase

Membrane-bound ATPase from E. coli can easily dissociate into

two parts: the FO or transmembrane proton translocating portion and

the ATPase active F1 portion (Vogel and Steinhart, 1976) with bound

annular lipids (Peter and Ahlers, 1975). Reconstitution experiments

were performed using solubilized F1 ATPase and ATPase depleted

membranes to further examine the importance of bound lipids versus

bulk membrane in ethanol resistance. Depleted membranes from


























(D O





cm a





eo





,aQ



CL~U



~o _


a,

CUr

00




ru2






















m








0o
o0
iy






O -S




















c,













(Jnno4 x u!aloid 6wlsalowoJcnw )
,J.AIIJ. aso-dLV









ethanol-grown cells were reconstituted with soluble F1 ATPase from

control cells. The break temperature of this preparation was increased

by the inclusion of ethanol (figure 10) indicating that the changes in

bulk membrane alone do not provide protection against ethanol.

Membranes from control cells were also reconstituted with F ATPase

from ethanol-grown cells. This reconstituted ATPase was resistant to

the effect of ethanol on break temperature (figure 10B) As a control,

ATPase and stripped membranes from ethanol-free cells were reconsti-

tuted. This preparation exhibited the same ethanol-induced increase in

break temperature as native membranes. Thus the sensitivity and

resistance of ATPase to the effects of ethanol on break temperature of

Arrhenius plots is a property of the soluble portion containing protein

and bound annular lipids, independent of the bulk membrane.

Cooperativity of ATPase Inhibition by Sodium

Membrane-bound ATPase of E. coli is inhibited by sodium ions with

Ca+ as a cofactor (Evans, 1969). This inhibition is cooperative with

a Hill coefficient (n) greater than one (Sinerez et al., 1973). Unlike

the break temperature observed in Arrhenius plots, the cooperativity of

this inhibition is strictly dependent upon the ATPase being membrane-

bound. Cooperativity of sodium inhibition is easily destroyed by

treatments such as freezing, assaying at a temperature above or below

the growth temperature or by solubilizing the enzyme (table IV).

Figure 11 shows the effect of ethanol on a Hill plot of ATPase.

Inclusion of ethanol in assays reduced the Hill coefficient. Dimethyl






























PLm
0










0 0U






L-r
Efl
Cd Cd
Cd~~












E r
:r "



Cd o








,"4 4
CdY=~-

Cd C Cf
S0 ~i~
m Cd
Cfl 0
Cd Cd3d

~ ~C
--C~ .g-.
C C
oI -la
oL

~Cfl~
Cd Cd C
0d~~ -, 0
Cda~

Cd C~r C
.Cd~a


U O




300m
Cd~0d
*Cd4.4"



















































































L -!
oJ


(Jnoq x uialojd bUJ/salowuolUOuj)


G3ZAo10aAH dIV


I I I


o o 0
N


0






0
0
o






0
" X




























0
0










0 0
xn













0
o0

Y*


if


I I I
0 0


I I


* 0















Table IV. Effect of a Variety of Physical Treatments on the
Cooperativity of Sodium Inhibition of ATPase from E. colia


Treatment

None

Freeze and thawed membranes

Solubilized ATPase

Assayed at 42C

Assayed at 300C


Hill Coefficient

1.26+ 0.08 (12)

1.02 (1)

1.03t 0.08 (3)

1.12 0.04 (2)

1.09 t 0.02 (2)


aE. coli TB-4 was grown at 370C and assayed at the
growth temperature unless otherwise specified.























Figure 11. Hill plots of the sodium inhibition of membrane-
bound ATPase. Assays were performed in the absence (*) and
the presence (0) of ethanol (16g/L). The sodium concentration
was varied over the range of 0-250nM.























1.0 h


Oh


In SODIUM CONCENTRATION


- i.0 o








sulfoxide, pentobarbital and chloropromazine also reduced the co-

operativity of sodium inhibition (table V). Cooperativity was not

reduced by ethanol in membrane-bound ATPase isolated from cells grown

in the presence of ethanol (table V) Additionally, ATPase from ethanol-

grown cells was resistant to dimethyl sulfoxide and pentobarbital but

remained sensitive to chloropromazine (table V). Pentobarbital (Ingram

et al., 1978) and dimethyl sulfoxide (Ingram, 1977) induce changes in

fatty acid composition similar to those caused by ethanol while

chloropromazine (Ingram et al., 1978) induces nearly opposite changes.

The protection provided against other agents by growth in the presence

of ethanol suggests that some adaptive change has occurred involving

lipids which compensates for the direct physical effect of these agents

on ATPase.

To investigate whether changes in bulk membrane fatty acid

composition per se are sufficient to afford protection against the direct

effect of ethanol on the cooperativity of sodium inhibition, the fatty

acid composition of cells was artificially altered. E. coli was grown

in the presence of exogenous oleic acid (18:1) to increase the proportion

of unsaturated fatty acids within the membrane (table I). Under these

conditions, no protection was provided against the effect of ethanol

(table VI) in spite of the increased proportion of unsaturated fatty acids.

Growth of E. coli with exogenous palmitic acid (16:0) decreased the

proportion of unsaturated fatty acids within the membrane (table I) and

also had no effect on ethanol sensitivity (table VI) Thus changes














Table V. The Effect of Ethanol and other Membrane Agents on
the Hill Coefficients of Membrane-bound ATPase


HILL COEFFICIENTa

Control Grown with
Assay Additive (n S. D.)b Ethanol (16g/L)
(n S. D.)a


None 1.26 + 0.08 (12) 1.28 0.06 (7)
Ethanol 16g/L 0.89 0.14 (9) 1.27 +0.09 (8)
Dimethyl Sulfoxide 33g/L 0.98 0.06 (4) 1.24 0.03 (2)
Pentabarbital 20g/L 0.93 + 0.03 (2) 1.26t 0.02 (2)
Chloropromazine 0.2g/L 1.03 0.05 (2) 1.01 0.04 (2)


aNumber within parentheses
of replicates.


to the right of S. D. denotes number


b. D., standard deviation.















I I
I I I IC *0


M C) C '4 00 o 0 (.0 *F 0


_ C 0 0 "

So
00 0000. . -

S000 0 0000 o


a) j +0 + I + + I +I +I +1 4-



> Ua 0 0 0
0 D c> 3 0 1 c*, m L 0 0 0.

S1 o .0 a
S2 I i c i 4 C ) ,
00 I -0 0 0 0 ;












U'0 CD 0o



o Q o. . .0 O. A
. Ioo 1o a
1z u Lo -

F0 I 0 0 3
So oo





C) 1 0 a
. g 5i 0 g i

CO I D S 0 C^ 3 C! o
0~0000C00 00











0 C-S 0C .0 0 ^ c>
2 u a j u ro 0 M .


gU~( 55 ( g
ccu - Q







45
in the bulk lipid composition are not sufficient to provide protection

against the effect of ethanol on the Hill coefficient.

The role of growth and new protein synthesis in the acquisition of

resistance to ethanol was investigated. Chloramphenicol was added

to a culture at a concentration sufficient to stop protein synthesis.

The culture was then split into two parts and incubation continued for

three hours. One part was supplemented with ethanol (16g/L) while the

other served as a control. Membranes were isolated from both of these

cultures for the determination of Hill coefficients in the presence and

absence of ethanol (table VI). ATPase from the control cells was

sensitive to the addition of ethanol while the cells which were exposed

to ethanol were resistant to the reduction in the Hill coefficient by

ethanol. This indicates that the acquisition of ethanol resistance does

not require new protein synthesis. In a similar experiment, a stationary

phase culture was split into two parts. Ethanol (16g/L) was added to

one part while the other served as a control. Following incubation for

an additional 16 hours at 370C, cells were harvested and the Hill

coefficients determined for each in the presence and absence of ethanol

(table VI). Stationary cells incubated in the absence of ethanol remained

sensitive while those incubated in the presence of ethanol acquired

resistance to ethanol. The bulk lipid composition of membranes in

stationary phase cells was not altered by the addition of ethanol (table I).

This provides further evidence that changes in bulk lipid composition

are not essential for the amelioration of ethanol effects.









In Vitro Acclimation of ATPase

Our previous experiments with Hill coefficients and Arrhenius plots

indicated that the adaption of membrane-bound ATPase to ethanol does

not require new protein synthesis or changes in the bulk lipid composi-

tion. This hypothesis was further examined in vitro. Isolated membranes

were incubated for 12 hours at 370 C in the presence and absence of

ethanol. The cooperativity of sodium inhibition was protected from the

ethanol effect following prior incubation in the presence of ethanol

(table VI), analogous to the results obtained in vivo with whole cells.

Likewise, the Arrhenius plots of the membrane-bound ATPase incubated

for 12 hours with ethanol were resistant to the inclusion of ethanol

(figure 12). Similar experiments, in vitro, were attempted with soluble

ATPase (7 & 12 hour incubations). Storage of the soluble ATPase in

ethanol, however, resulted in Arrhenius plots which did not exhibit a

distinct discontinuity in their slopes. The adaptation of membrane-

bound ATPase in vitro indicates that resistance can be acquired by a

rearrangement of ATPase with existing membrane components. The

failure of soluble ATPase to maintain its native thermodynamic

properties following prolonged incubation in the presence of ethanol

suggests that this rearrangement may involve other components in

addition to the bound lipids.




















Figure 12. Arrhenius plots of membrane-bound ATPase
previously exposed to ethanol (16g/L) in vitro. Assays
were performed in the absence (e) and presence (0) of
16g/L of ethanol.



















20.0-



0
I 0.0-
>x


O a
<. 5.0 -


Q.--
0
< E

E








1I I .

3.2 3.4 3.5 3.6
K 000
I/ K x 1000












DISCUSSION

Our results indicate that growth in the presence of ethanol results

in changes which confer resistance to the effects of ethanol on two

membrane-bound enzymes. The lac permease is inhibited in vivo by

the initial addition of ethanol to cells grown without ethanol. During

growth in the presence of ethanol, this inhibition is ameliorated (figure

2). Membrane-bound ATPase (Mg++) from cells grown without ethanol

is inhibited in vitro by the addition of ethanol at 150 C and stimulated

by the addition of ethanol at 370 C (figures 5 & 6). Again ATPase

preparations from cells grown in the presence of ethanol are substantially

resistant to both effects. Arrhenius plots of lac permease and ATPase

exhibit a discontinuity, the position of which is shifted to an elevated

temperature by the inclusion of ethanol in the assay mixture (figures 7

& 8). Enzyme preparations from cells grown in the presence of ethanol

are resistant to this effect of ethanol. Ethanol eliminates the

cooperativity of Na+ inhibition of ATPase from control cells. Again,

preparations from cells grown in the presence of ethanol are resistant

to ethanol. Thus adaptive changes occur during growth in the presence

of ethanol which ameliorate the direct effect of ethanol on these

membrane functions.

The effect of alcohols on biological systems appears to result from

their interaction at hydrophobic sites (Ingram, 1976; and Seeman, 1972).

49





50

These hydrophobic sites need not be limited to the membrane lipid bilayer

but may also include the hydrophobic regions of proteins and their

annular lipids. Following prolonged exposure to ethanol two types of

membrane adaptation are frequently observed. The first involves changes

in physical properties of membrane-bound enzymes (Hosein et al. 1977;

and Israel and Kuriyama, 1971) and the second involves changes in the

bulk lipid composition of membranes (Ingram, 1976; Littleton and John,

1977; and Miceli and Ferrel, 1973).

Membrane-bound enzymes are affected by changes in their lipid

environment (Esfahani et al., 1971; Kimelberg and Papahadjopoulas,

1972; and Mavis and Vagelos, 1972). The interaction of drugs like

ethanol with the membrane alters the physical properties of the membrane

and affects the activities of membrane-bound enzymes (Lee, 1976).

The changes in bulk lipid composition following growth in the presence

of ethanol (table I) have been postulated as part of an adaptive response

to compensate for some of these effects (Ingram, 1976).

We have examined the role of the lipids in the adaptation of

membrane-bound enzymes. Our results with membrane-bound and

soluble ATPase (table III) indicates that the effect of ethanol on

Arrhenius plots of this enzyme is mediated through a direct interaction

of ethanol with the enzyme and perhaps annular lipid rather than through

changes in bulk membrane. Reconstitution experiments indicate that

sensitivity or resistance to ethanol are determined within the solubilized

portion containing bound lipids, independent of bulk membranes. The







51

adaptation of ATPase to ethanol in E. coli strain WN I (figure 9) as

well as the adaptation of stationary cells (table VI) and isolated

membranes (figure 12 and table VI) to ethanol indicates that the bulk

lipid changes are not essential for the adaptation of membrane-bound

enzymes to ethanol. However, changes in lipid composition may be of

significance for other cellular function in vivo.

Membrane-bound enzymes have been shown to exhibit some

selectivity for particular phospholipids from their environment which

allows maximal enzymatic activity (Warren et al., 1975). Indeed, the

isolation of different regions of the membrane has provided direct

physical evidence that membrane proteins may partition between specific

lipid domains in vivo (van Heerikhuisen et al., 1975; and Letellier et

al., 1977). These associations with particular annular lipids presumably

maintain the enzyme in the most thermodynamically favored configuration.

Thus membrane-bound enzymes appear to behave somewhat like a

cartesian diver in that they appear to select the proper environment

(i. e. annular lipid composition) to maintain the thermodynamically

favored state. The intercalation of ethanol results in changes in the

membrane environment. In the presence of ethanol, membrane-bound

enzymes may exchange their annular lipids for more unsaturated species

to maintain the most thermodynamically favored state.

This cartesian diver model explains how membrane-bound enzymes

could adapt in vitro to ethanol. The selection of a new environment by

membrane-bound enzymes would only require the proper unbound lipids








within the membrane with which the enzyme could reassociate. In

this model, the increase in unsaturated fatty acids following growth

in the presence of ethanol could maintain the proper microviscosity in

the bulk lipid bilayer which would have become more saturated in

nature due to the removal of unsaturated phospholipid species by this

protein/lipid reassociation. Alternatively, an increase in unsaturation

of molecular species could increase the proportion of the lipid environ-

ment with the physical properties which optimize enzyme function.

This model suggests that cells with naturally higher levels of unsatu-

rated fatty acids in their bulk lipids would be capable of coping with

ethanol better than cells with lower levels of unsaturated fatty acids.

Recently we have isolated fifteen mutants of E. coli which are resistant

to ethanol all of which contain higher proportions of unsaturated fatty

acids than the parental strain when grown in the absence of ethanol.

This model for the development of ethanol tolerance for membrane-

bound enzymes may be related to the observed consequence of prolonged

alcohol consumption. An initial acute tolerance occurs following

several hours of exposure to ethanol. This tolerance may be due to

rearrangement of membrane proteins with existing lipids as described

in our model. Chronic tolerance and its associated dependence could

be due to both this rearrangement and the ethanol-induced changes in

membrane composition (Ingram et al., 1978; and Littleton and John,

1977). These lipid changes need not be limited to phospholipids and






53

fatty acids. Other membrane components such as cholesterol and

carotenoids may also be involved.














BIBLIOGRAPHY


Buttke, T. M., and Ingram, L. 0. (1978), Biochemistry 17:637-644.

Cantley, L. C. Jr., Josephson, L., Warner, R., Yanagisawa, M.,
Lechene, C., and Guidotti, G. (1977), L. Biol.Chem.252:
7421-7423.

Chang, T., Lewis, J., and Glazko, A. J. (1967), Biochim. Biophys.
Acta 135:1000-1007.

Cronan, J. E. Jr. (1975), i. Biol. Chem. 250:7074-7077.

Evans, D. J. Jr. (1969), J. Bacteriol. 114:239-248.

Esfahani, M., Lunbreck, A. R., Knutton, S. K., Oka, T., and Wakil,
S. J. (1971), Proc. Natl. Acad. Sci. USA 68:3180-3184.

Fox, J. E., Bourdages, R., and Beck, I. T. (1978), AM. J. Dig. Dis.
23:193-200.

Fried, V. A., and Novick, A. (1973), J. Bacteriol. 114:239-248.

Fulco, A. J. (1974), Ann. Rev. Biochem. 43:215-241.

Futai, M., Sternweis, P. C., and Heppel, L. A. (1974), Proc. Natl.
Acad. Sci. USA 71:2725-2729.

Gelmann, E. P., and Cronan, J. E. Jr. (1972), J. Bacteriol. 112:
381-387.

Grisham, C. M., and Barnett, R. E. (1972), Biochim. Biophvs. Acta
266:613-624.

van Heerikhuisen, H., Kwah, E., van Brugzen, E. E. J. and Wilhott,
B. (1975), Biochim. Biophys. Acta 413:177-191.

Hesketh, T. R., Smith, G. A., Houslay, M. D., McGill, K. A.,
Birdsall, N. J. M., Metcalfe, J. C., and Warren, G. B. (1976),
Biochemistry 15:4145-4151.






55

Hill, M. W., and Bangham, A. D. (1975), In M. M. Gross (Ed.),
Alcohol Intoxication and Withdrawal. Experimental Studies. II,
pp 1-9, New York, Plenum Press.

Hosein, E. A., Hoffmann, I., and Linder, E. (1977), Arch. Biochem.
Biophys. 183:64-72.

Hoyumpa, A. M., Nicols, S. G., Wilson, F. A., and Schenker, S.
(1977), Lab.Clin. Med. 90:1086-1095.

Ingram, L. 0. (1977), A-o. Env. Microbiol. 33:1233-1236.

Ingram, L. 0. (1976), J. Bacteriol. 125:670-678.

Ingram, L. O., Ley, K. D., and Hoffmann, E. M. (1978), Life Sciences
22:489-494.

Israel, M., and Kuriyama, K. (1971), Life Sciences 10:591-599.

Israel, Y., Salazar, I., and Rosenmann, E. (1968), J. Nutr. 96:499-504.

Kanfer, I., and Kennedy, E. P. (1963),J. Biol. Chem. 238:2919-2922.

Kepes, A. (1971), J. Memb. Biol. 4:87-112.

Kimelberg, H. K., and Papahadjopoulas, D. (1972), Biochim. Biophvs.
Acta 282:277-292.

Lee, A. G. (1977), Mol. Pharmacol. 13:474-485.

Lee, A. G. (1976), Nature 262:545-548.

Lenaz, G., Parenti-Castelli, G., and Sechi, A. M. (1975), Arch.
Biochem. Biophys. 167:72-79.

Letellier, L., Moudden, H., and Shechter, E. (1977), Proc. Natl.
Acad. Sci. USA 74:452-456.

Linden, C. D., Wright, K. L., McConnell, H. M., and Fox, C. F.
(1973), Proc. Natl. Acad. Sci. USA 70:2271-2275.

Littletcn, J. M., and John, G. (1977),J. Pharm. Pharmac. 29:579-580.

Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J.
(1951), J. Biol. Chem. 193:265-275.








Luria, S. E., and Delbruck, M. (1943), Genetics 28:491-511.

Massey, V., Curti, B., and Ganther, H. (1966), J. Biol. Chem.
241:2347-2357.

Mavis, R. D., and Vagelos, P.R. (1972), J. Biol. Chem. 247:652-659.

Miceli, J. M., and Ferrel, W. J. (1973), Lipids 8:722-727.

Miller, J. H. (1972) Experiments in Molecular Genetics, pp. 301,
Cold Spring Harbor Laboratory, Cold Spring Harbor, N. Y.

Mitjavila, S., Lacombe, C., and Carrera, G. (1976), Biochem.
Pharmacol. 25:625-630.

Morrisett, J. D., Pownall, H. I., Plumlee, R. T., Smith, L. C.,
Zehner, Z., Esfahani, M., and Wakil, S. J. (1975), i. Biol.
Chem. 250:6969-6976.

Overath, P., Hill, F. F., and Lannek-Hirsch, I. (1971), Nature
New Biology 234:264-267.

Peter, H. W., and Ahlers, J. (1974), Arch. Biochem. Biophys.
170:169-178.

Post, R. L., Hegyvary, C., and Kume, S. (1972),J. Biol. Chem.
247:6530-6540.

Putman, S., and Koch, A. L. (1975), Anal. Biochem. 63:350-360.

Rathburn, W. B., and Betlach, M. V. (1969), Anal. Biochem. 28:436-445.

Rosen, B. P., and Adler, L. A. (1975), Biochim. Biophys. Acta
387:23-26.

Seeman, P. (1972), Pharmacol. Rev. 24:583-655.

Sinensky, M. (1974), Proc. Natl. Acad. Sci. USA 71:522-525.

Sinerez, F., Farias, R. N., and Trucco, R. E. (1973), FEBS Letters
32:30-32.

Sullivan, K. H., Jain, M. K., and Koch, A. L. (1974), Biochim.
Biophys. Acta 352:287-297.







57

Sun, A. Y. (1976), Ann. N. Y. Acad. Sci. 273:295-302.

Thilo, L., Trauble, H., and Overath, P. (1977), Biochemistry
16:1283-1290.

Vogel, G., and Steinhart, R. (1976), Biochemistry 15:208-216.

Warren, G. B., Houslay, M. D., Birdsall, N. J. M., and Metcalfe,
J. C. (1975), Nature (London) 255:684-687.

Worthington, B. S., Meserole, L., and Syrotuck, J. A. (1978),
Am. I. Dig. Dis. 23:23-32.














BIOGRAPHICAL SKETCH

Benjamin Fisher Dickens Jr. was born on August 1, 1951, in

Jacksonville, Florida. He lived in Fernandina Beach, Florida, with

his parents, Dr. and Mrs. B. F. Dickens, from 1951 until 1969 when

he graduated from Fernandina Beach Senior High School. In September

1969 he began attending college at the University of Florida in

Gainesville, Florida. He obtained his Bachelor of Science in June

1973 and in March 1976 he obtained his Master of Science from the

University of Florida. He married Bonnie Estelle Quattlebaum on July

1, 1972, and their son, Brian Fisher, was born on April 21, 1975.

Benjamin is a candidate for the degree of Doctor of Philosophy in

December 1978.









I certify that I have read this study and that in my opinion It
conforms to acceptable standards of scholarly presentation and is
fully adequate, in scope and quality, as a dissertation for the degree
of Doctor of Philosophy.



Lonnie O. Ingram, Chairm n
Associate Professor of Microbiology

I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is
fully adequate, in scope and quality, as a dissertation for the degree
of Doctor of Philosophy.



Edward M. Hoffmanr /i
Associate Professor of Microbiology

I certify that I have read this study and that in my opinion it
conforms to acceptable standards of scholarly presentation and is
fully adequate, in scope and quality, as a dissertation for the degree
of Doctor of Philosophy.




Charles M. Allen Jr.
Associate Professor of Biochemistry


This dissertation was submitted to the Graduate Faculty of the Depart-
ment of Microbiology and Cell Science in the College of Liberal Arts and
Sciences and to the Graduate council, and was accepted as partial
fulfillment of the requirements for the degree of Doctor of Philosophy.

December 1978




Dean, Graduate School